Molecular and Diagnostic Procedures in Mycoplasmology Volume II
DIAGNOSTIC PROCEDURES
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Molecular and Diagnostic Procedures in Mycoplasmology Volume II
DIAGNOSTIC PROCEDURES Edited by
JOSEPH G. TULLY Mycoplasma Section Laboratory of Molecular Microbiology National Institute of Allergy and Infectious Diseases Frederick Cancer Research and Development Center Frederick, Maryland
SHMUEL RAZIN Department of Membrane and Ultrastructure Research The hiebrew University-Hadassah Medical School Jerusalem, Israel
ACADEMIC PRESS San Diego
New York
Boston
London
Sydney
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Toronto
Front cover photograph: Vero cells infected with M. hyorhinis, stained by a double stain method using DNAF and fluoresceinated anti-M hyorhinis antibodies, and viewed with the filter set for fluorescein. Courtesy of Dr. Gerald K. Masover and Frances A. Becker, Genentech, Inc., South San Francisco, CA.
This book is printed on acid-free paper. ©
Copyright © 1996 by ACADEMIC PRESS, INC. All Rights Reserved. No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher.
Academic Press, Inc. A Division of Harcourt Brace & Company 525 B Street, Suite 1900, San Diego, California 92101-4495 United Kingdom Edition published by Academic Press Limited 24-28 Oval Road, London NWl 7DX Library of Congress Cataloging-in-Publication Data Molecular and diagnostic procedures in mycoplasmology / edited by Shmuel Razin, Joseph G. Tully. p. cm. Includes indexes. Contents: v. 1. Molecular characterization - v. 2. Diagnostic procedures. ISBN 0-12-583805-0 (v. 1: alk. paper) ISBN 0-12-583806-9 (v. 2: alk. paper) 1. Mycoplasma diseases-Diagnosis. 2. Mycoplasma diseases-Molecular aspects. I. Razin, Shmuel. II. Tully, Joseph G. [DNLM: 1. Mycoplasma—physiology. 2. Mycoplasma—pathogenicity. 3, Molecular Biology-methods. 4. Mycoplasma Infections-diagnosis. QW 143 M718 1995J QR201.M97M63 1995 589.9-dc20 DNLM/DLC for Library of Congress 95-4586 CIP
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Contents
Contributors
xi
Preface
xv
Contents of Volunne I
xvii
Mollicute-Host Interrelationships: Current Concepts and Diagnostic Implications Joseph C. Tully
SECTION A Diagnostic Genetic Probes A1
Introductory Remarks
25
Shmuel Razin A2
Oligonucleotide Probes Complementary to 16S rRNA
29
Karl-Erik Johansson A3
Cloned Genomic DNA Fragments as Probes
47
David Yogev and Shmuel Razin A4
PCR: Selection of Target Sequences
53
Remi Kovacic, Odile Crau, and Alain Blanchard A5
PCR: Preparation of DNA from Clinical Specimens
61
Bertille de Barbeyrac, Christiane Bebear, and David Taylor-Robinson A6
PCR: Amplification and Identification of Products Bertille de Barbeyrac and Christiane Bebear
65
VI A7
Contents PCR: Application of Nested PCR to Detection of Mycoplasmas
75
Ryo Harasawa A8
PCR: Random Amplified Polymorphic D N A Fingerprinting
81
Steven I. Geary and Mark H. Forsyth
SECTION B Bl
Immunological Tools
Introductory Remarks
89
Joseph C. Tully B2
ELISA in Small Animal Hosts, Rodents, and Birds
93
M. B. Brown, j. M. Bradbury, and I. K. Davis B3
ELISA in Large Animals
105
I. Nicolet and I. L. Martel B4
ELISA in Human Urogenital Infections and AIDS
115
Richard Yuan-Hu Wang and Shyh-Ching Lo B5
ELISA in Respiratory Infections of Humans
123
Gail H. Cassell, Ginger Gambill, and Lynn Duffy B6
Monoclonal Antibodies as Diagnostic Tools
137
Ghester B. Thomas, Monique Gamier, and John T. Boothby B7
Microimmunofluorescence David
B8
147
Taylor-Robinson
Immunoblots and Immunobinding
151
David Thirkell and Bernard L Precious B9
Differentiation of Mycoplasma Mycoplasma
pneumoniae
Joseph G. Tully
genitalium
from
by Immunofluorescence
169
Contents
SECTION C Cl
vii
Antibiotic Sensitivity Testing
Introductory Remarks
181
Christiane Bebear C2
Problems and Opportunities in Susceptibility Testing ofMoliicutes
185
George E. Kenny C3
Determination of M i n i m a l Inhibitory Concentration
189
Christiane Bebear and Janet A. Robertson C4
Cidal Activity Testing David
Taylor-Robinson
SECTION D D1
199
Diagnosis of Specific Diseases
Introductory Remarks
207
Joseph C. Tully D2
Laboratory Diagnosis of Mycoplasma R. J. Harris, j. Williamson,
D3
Infection
211
C. Hahn, and B. P. Marmion
Diagnosis of Sexually Transmitted Diseases David
D4
pneumoniae
225
Taylor-Robinson
Diagnosis of Neonatal Infections
237
Ken B. Waites and Gail H. Cassell D5
Mycoplasmas in AIDS Patients
247
Shyh-Ching Lo D6
Mycoplasma Infections of Cattle Ed A. ter Laak and H. Louise Ruhnke
255
VIM D7
Contents Mycoplasma Infections of Goats and Sheep
265
A, I. DaMassa D8
Mycoplasma Infections of Swine
275
Richard F. Ross and Gerald W. Stemke D9
Mycoplasma Infections of Poultry
283
Stanley H, Kleven and Sharon Levisohn DIO
Diagnosis of Spiroplasma Infections in Plants and Insects
293
C. Saillard, C. Barthe, ]. M. Bove, and R. F. Whitcomb Dll
Detection of Phytoplasma Infections in Plants
299
£. Seemueller and B. C. Kirkpatrick D12
Identification of Mollicutes from Insects
313
Robert F. Whitcomb and Kevin /. Hackett
SECTION E Experimental Infections El
Introductory Remarks
325
Joseph G. Tully E2
Experimental Mycoplasmal Respiratory Infections in Rodents
327
Gail H. Cassell and A. Yancey E3
Urogenital Infections in Rodents Patricia M. Furr and David
E4
337
Taylor-Robinson
Experimental Models of Arthritis
349
Leigh Rice Washburn E5
Experimental Infections in Poultry Janet M. Bradbury and Sharon Levisohn
361
Contents E6
Experimental Infections of Swine
ix 371
Marylene Kobisch and Richard F. Ross E7
Experimental Infections in Cattle
377
Ricardo F. Rosenbusch and H. Louise Ruhnke E8
Experimental Infections of Plants by Spiroplasmas
385
X. Foissac, I. L Danet, C. Saillard, R. F. Wbitcomb, and I. M. Bove E9
Experimental Phytoplasma Infections in Plants and Insects
391
Alexander H. Purcell ElO
Mycoplasmas and in Vitro Infections of Cell Cultures w i t h HIV Shyb-Ching Lo and Alain
SECTION F
Fl
399
Blancbard
Diagnosis of Mycoplasma Infections of Cell Cultures
Introductory Remarks
407
losepb G. Tally F2
Isolation of Mycoplasmas from Cell Cultures by Axenic Cultivation Techniques
411
Ricbard A. Del Giudice and losepb G. Tully F3
Detection of Mycoplasmas by D N A Staining and Fluorescent Antibody Methodology
419
Gerald K. Masover and Frances A. Becker F4
Detection of Mycoplasma Infection by PCR
431
Connie Veilleux, Sbmuel Razin, and Laurie H. May F5
Antibiotic Treatment of Mycoplasma-lnfected Cell Cultures Ricbard A. Del Giudice and Roberta S. Gardella
439
X
Contents
F6
Prevention and Control of Mycoplasma Infection of Cell Cultures
445
Ann Smith and Ion Mowles
APPENDIX Table I
Genus Mycoplasma and Major Characteristics
455
Table II
Genera Ureaplasma and Acholeplasma
458
Table III
Genera Anaeroplasma and Asteroleplasma
458
Table IV
Genera Entomoplasma and Mesoplasma
459
Table V
Group Classification of Genus Spiroplasma
460
Index
463
Contributors
Numbers in parentheses indicate tiie pages on which the authors' contributions begin.
C. Barthe (293), Laboratoire de Biologic Cellulaire et Moleculaire, Institut National de Recherche Agronomique, F-33883 Villenave d'Ornon, France Christiane Behear (61, 65, 181, 189), Laboratoire de Bacteriologie, Universite de Bordeaux II, F-33076 Bordeaux, France Frances A. Becker (419), Department of Quality Control, Genentech, Inc., South San Francisco, California 94080 Alain Blanchard (53, 399), Department du SID A et des Retrovirus, Institut Pasteur, Unite d'Oncologie Virale, 75724 Paris, France John T. Boothby (137), Department of Biological Sciences, San Jose State University, San Jose, California 95192 J. M. Bove (293, 385), Laboratoire de Biologic Cellulaire et Moleculaire, Institut National de Recherche Agronomique, F-33883 Villenave d'Ornon, France Janet M. Bradbury (93, 361), Department of Veterinary Pathology, University of Liverpool, Leahurst, Neston, South Wirral L64 7TE, United Kingdom M. B. Brown (93), Department of Infectious Diseases, College of Veterinary Medicine, University of Florida, Gainesville, Florida 32611 Gail H. Cassell (123, 237, 327), Department of Microbiology, University of Alabama at Birmingham, Birmingham, Alabama 35294 A. J. DaMassa (265), Department of Population Health and Reproduction, School of Veterinary Medicine, University of California at Davis, Davis, California 95616 J. L. Danet (385), Laboratoire de Biologic Cellulaire et Moleculaire, Institut National de Recherche Agronomique, F-33883 Villenave d'Ornon, France J. K. Davis (93), Department of Comparative Medicine, University of Florida, Gainesville, Florida 32611 Bertille de Barbeyrac (61, 65), Laboratoire de Bacteriologie, Universite de Bordeaux II, F-33076 Bordeaux, France Richard A. Del Giudice (411, 439), Mycoplasma Laboratory, SAIC Frederick, NCI-Frederick Cancer Research and Development Center, Frederick, Maryland 21702 Lynn Duffy (123), Department of Microbiology, University of Alabama at Birmingham, Birmingham, Alabama 35294 X. Foissac (385), Laboratoire de Biologic Cellulaire et Moleculaire, Institut National de Recherche Agronomique, F-33883 Villenave d'Ornon, France Mark H. Forsyth (81), Department of Pathobiology, University of Connecticut, Storrs, Connecticut 06269 Patricia M. Furr (337), MRC Sexually Transmitted Diseases Research Group, Department of Genitourinary Medicine, St. Mary's Hospital Medical School, London W2 INY, United Kingdom Ginger Gambill (123), Department of Microbiology, University of Alabama at Birmingham, Birmingham, Alabama 35294
XI i
Contributors
Roberta S. Gardella (439), Mycoplasma Laboratory, SAIC Frederick, NCI-Frederick Cancer Research and Development Center, Frederick, Maryland 21702 Monique Gamier (137), Laboratoire de Biologic Cellulaire et Moleculaire, Institut National de Recherche Agronomique, F-33883 Villenave d'Ornon, France Steven J. Geary (81), Department of Pathobiology, University of Connecticut, Storrs, Connecticut 06269 Odile Grau (53), Department du SIDA et des Retrovirus, Institut Pasteur, Unite d'Oncologie Virale, 75724 Paris, France Kevin J. Hackett (313), Insect Biocontrol Laboratory, Plant Sciences Institute, United States Department of Agriculture, Agricultural Research Service, BARC, Beltsville, Maryland 20705 C. Hahn (211), Department of Biochemistry, University of Adelaide, Adelaide, South Australia 5000, Australia Ryo Harasawa (75), Animal Center for Biomedical Research, Faculty of Medicine, University of Tokyo, Hongo, Tokyo 113, Japan R. J. Harris (211), School of Pharmacy and Medical Sciences, University of South Australia, Adelaide, South Australia 5000, Australia Karl-Erik Johansson (29), Laboratory of Bacteriology, Research and Development, The National Veterinary Institute, S-750 07 Uppsala, Sweden George E. Kenny (185), Department of Pathobiology, School of Public Health and Community Medicine, University of Washington, Seattle, Washington 98195 B. C. Kirkpatrick (299), Department of Plant Pathology, University of California at Davis, Davis, California 95616 Stanley H. Kleven (283), Department of Avian Medicine, University of Georgia School of Veterinary Medicine, Athens, Georgia 30602 Marylene Kobisch (371), Centre National d'Etudes Veterinaires et Alimentaires, Station de Pathologic Porcine, F-22440 Ploufragan, France Remi Kovacic (53), Department du SIDA et des Retrovirus, Institut Pasteur, Unite d'Oncologie Virale, 75724 Paris, France Sharon Levisohn (283, 361), Department of Poultry Diseases, Ministry of Agriculture Veterinary Services and Animal Health, Kimron Veterinary Institute, Beit-Dagan 50250, Israel Shyh-Ching Lo (115, 247, 399), Department of Infectious and Parasitic Disease Pathology, Division of Molecular Pathology, American Registry of Pathology, Armed Forces Institute of Pathology, Washington, District of Columbia 20306 B. P. Marmion (211), Department of Pathology, Institute of Medical and Veterinary Science, University of Adelaide, Adelaide, South Australia 5000, Australia J. L. Martel (105), Laboratoire de Reference de I'OIE pour la PPCB, CNEVA-LPB, F-69342 Lyon, France Gerald K. Masover (419), Department of Quality Control, Genentech, Inc., South San Francisco, California 94080 Laurie H. May (431), Genentech, Inc., South San Francisco, California 94080 Jon Mowles (445), Biochem ImmunoSystems Ltd., Woking GU21 5JY, United Kingdom J. Nicolet (105), Institute of Veterinary Bacteriology, University of Berne, CH-3001 Berne, Switzerland
Contributors
xiii
Bernard L. Precious (151), Division of Cell and Molecular Biology, School of Biological and Medical Sciences, University of St. Andrews, St. Andrews, Fife KY16 9AL, Scotland Alexander H. Purcell (391), Department of Environmental Science, Policy, and Management, University of California at Berkeley, Berkeley, California 94720 Shmuel Razin (25, 47, 431), Department of Membrane and Ultrastructure Research, The Hebrew University-Hadassah Medical School, Jerusalem 91120, Israel Janet A. Robertson (189), Department of Medical Microbiology and Infectious Diseases, University of Alberta, Edmonton, Alberta, Canada T6G 2H7 R. F. Rosenbusch (311), Veterinary Medical Research Institute, College of Veterinary Medicine, Iowa State University, Ames, Iowa 50011 Richard F. Ross (275, 371), College of Veterinary Medicine, Iowa State University, Ames, Iowa 50011 H. Louise Ruhnke (255, 377), Mycoplasma Laboratory, Veterinary Laboratory Services, Ontario Ministry of Agriculture and Food, Guelph, Ontario, Canada NIH 6R8 C. Saillard (293, 385), Laboratoire de Biologic Cellulaire et Moleculaire, Institut National de Recherche Agronomique, F-33883 Villenave d'Ornon, France E. Seemueller (299), Biologische Bundesanstalt fur Land- und Forstwirtschaft, Institut fur Pflanzenschutz im Obstbau, D-69216 Dossenheim, Germany Ann Smith (445), Cantab Pharmaceutical and Research Ltd., Cambridge CB4 4GN, England Gerald W. Stemke (275), Department of Biological Sciences, University of Alberta, Edmonton, Alberta, Canada T6G 2E9 David Taylor-Robinson (61, 147, 199, 225, 337), MRC Sexually Transmitted Diseases Research Group, Department of Genitourinary Medicine, St. Mary's Hospital Medical School, Paddington, London W2 INY, United Kingdom Ed A. ter Laak (255), Department of Bacteriology, DLO Institute for Animal Science and Health, 8200 AB Lelystad, The Netherlands David Thirkell (151), Division of Cell and Molecular Biology, School of Biological and Medical Sciences, University of St. Andrews, St. Andrews, Fife KYI6 9AL, Scotland Chester B. Thomas (137), Department of Pathobiological Sciences, School of Veterinary Medicine, University of Wisconsin, Madison, Wisconsin 53706 Joseph G. Tully (1, 89, 169, 207, 325, 407, 411), Mycoplasma Section, Laboratory of Molecular Microbiology, National Institute of Allergy and Infectious Diseases, Frederick Cancer Research and Development Center, Frederick, Maryland 21702 Connie Veilleux (431), Genentech, Inc., South San Francisco, California 94080 Ken B. Waites (237), Departments of Pathology, Microbiology, and Rehabilitation Medicine, Division of Laboratory Medicine, University of Alabama at Birmingham, Birmingham, Alabama 35233 Leigh Rice Washburn (349), Department of Microbiology, University of South Dakota School of Medicine, Vermillion, South Dakota 57069 R. F. Whitcomb (293, 313, 385), Insect Biocontrol Laboratory, Plant Sciences Institute, United States Department of Agriculture, Agricultural Research Service, BARC, Beltsville, Maryland 20705 J. Williamson (211), Department of Microbiology, Royal Hobart Hospital, Hobart, Tasmania 7000, Australia
XIV
Contributors
A. Yancey (327), Department of Microbiology, University of Alabama at Birmingham, Birmingham, Alabama 35294 David Yogev (47), Department of Membrane and Ultrastructure Research, The Hebrew University-Hadassah Medical School, Jerusalem 91120, Israel Richard Yuan-Hu Wang (115), Department of Infectious and Parasitic Disease Pathology, Division of Molecular Pathology, American Registry of Pathology, Armed Forces Institute of Pathology, Washington, District of Columbia 20306
Preface
The two volumes of Methods in Mycoplasmology published by Academic Press in 1983 have gained wide recognition as the most comprehensive and authoritative treatise on mycoplasma methodology, and are highly cited in the mycoplasma literature. These volumes have provided researchers and laboratory workers with well-tried and standardized procedures for the recovery, identification, and characterization of mycoplasmas. The developments in mycoplasmology which have taken place since the publication of these volumes have been outstanding due mainly to the application of molecular genetic methodology to mycoplasmas. Introduction of this methodology has had a significant impact on our understanding of the cell structure, genetics, metabolism, taxonomy, and phylogeny of mycoplasmas, as well as of the mechanisms of pathogenicity and the interaction of mycoplasmas with the immune system. These advances have found expression in the development of new diagnostic procedures, including those based on DNA probes and DNA amplification. As could be expected, significant developments have also taken place in the more "classical" procedures, those dealing with the cultivation, serological characterization, and pathogenicity testing of mycoplasmas. The two volumes of Molecular and Diagnostic Procedures in Mycoplasmology focus on the new procedures developed during the past decade, particularly those based on the new molecular methodology. This volume deals with the new genetic and immunological tools applied to the diagnosis of mycoplasma infections of humans, animals, plants, insects, and cell cultures. Volume I outlines the approaches, techniques, and procedures applied to cell and molecular biology studies of mycoplasmas. We are well aware that techniques outlined for rapidly moving subdisciplines may soon become dated. Yet experience gained through the use of the Methods in Mycoplasmology volumes confirms that the majority of methods detailed in the new volumes will continue to be useful for years to come. We thank our colleagues who were most helpful at the initial stages of selecting the topics and procedures to be covered and in developing the volume outlines. Considering the large number of chapters and contributors, keeping to the deadlines set by the publisher can be considered an outstanding achievement. Obviously, this could not have been accomplished without the cooperation of the many contributors. We express our gratitude and appreciation for their friendly cooperation in this endeavor. Shmuel Razin Joseph G. Tully
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Contents of Volume I Molecular Characterization
Molecular Properties of Moliicutes: A Synopsis Shmuel Razin SECTION
A
CULTIVATION AND MORPHOLOGY A1 Introductory Remarks Joseph C. Tully A2 Culture Medium Formulation for Primary Isolation and Maintenance of Moliicutes Joseph G. Tully A3 Cultivation of Spiroplasmas in Undefined and Defined Media Kevin J. Hackett and Robert F. Whitcomb A4 Insect Cell Culture Approaches in Cultivating Spiroplasmas Kevin j. Hackett and Dwight E. Lynn A5 Measurement of Mollicute Growth by ATP-Dependent Luminometry Janet A. Robertson and Gerald W. Stemke A6 Intracellular Location of Mycoplasmas David Taylor-Robinson A7 Localization of Mycoplasmas in Tissues Douglas j. Wear and Shyh-Ching Lo A8 Localization of Antigens on Mycoplasma Cell Surface and Tip Structures Duncan C. Krause and Maria K. Stevens
SECTION
B
GENOME CHARACTERIZATION AND GENETICS B1 Introductory Remarks Shmuel Razin B2 Isolation of Mycoplasma-like Organism DMA from Plant and Insect Hosts Bruce C. Kirkpatrick, Nigel A. Harrison, Ing-Ming Lee, Harold Neimark, and Barbara B. Sears B3 Mollicute Chromosome Size Determination and Characterization of Chromosomes from Uncultured Moliicutes Harold Neimark and Patricia Garle
Contents of Volume I
XVIII
B4 Physical and Genetic Mapping Thomas Proft and Richard Herrmann B5 Characterization of Virus Genomes and Extrachromosomal Elements Kevin Dybvig B6 Plasmid and Viral Vectors for Gene Cloning and Expression in Spiroplasma citri ). Renaudin and J. M. Bove B7 Artificial Transformation of Mollicutes via Polyethylene Glycol- and Electroporation-Mediated Methods Kevin Dybvig, Gail E. Gasparich, and Kendall W. King B8 DNA Methylation Analysis Aharon Razin and Paul Renbaum B9 Identification and Characterization of Genome Rearrangements Bindu Bhugra and Kevin Dybvig B10 Expression of Mycoplasmal Genes in Escherichia coli Paul Renbaum and Aharon Razin
SECTION C
MEMBRANE CHARACTERIZATION CI C2
C3 C4 C5 C6
SECTION D
Introductory Remarks Shmuel Razin Posttranslational Modification of Membrane Proteins Ake Wieslander, Susanne Nystrom, and Anders Dahlqvist Variant Membrane Proteins Kim S. Wise, Mary F. Kim, and Robyn Watson-McKown Membrane Fusion Shiomo Rottem and Mark Tarshis Mycoplasma Membrane Potentials Ulrich Schummer and Hans Gerd Schiefer Ion Flow and Cell Volume Shiomo Rottem
CELL METABOLISM D1 Introductory Remarks Shmuel Razin D2 Methods for Testing Metabolic Activities in Mollicutes j. Dennis Pollack
Contents of Volume I D3 Rapid Microcalorimetric and Ele troanalytical Measurements of Metabolic Activities R. j. Miles D4 Characterization of Heat Shock Proteins Christopher C. Dascher and jack Maniloff D5 Nucleolytic Activities of Mycoplasmas F. Chris Minion and Karalee j. jarvill-Taylor D6 Proteolytic Activities Tsuguo Watanabe and Ken-ichiro Shibata D7 Phospholipase Activity in Mycoplasmas Shiomo Rottem and Michael Salman
SECTION E
TAXONOMY AND PHYLOGENY El Introductory Remarks Shmuel Razin E2 Minimal Standards for Description of New Species of the Class Mollicutes Joseph C. Tully and Robert F. Whitcomb E3 Ribosomal RNA Sequencing and Construction of Mycoplasma Phytogenies Williann C. Weisburg E4 Restriction Endonuclease Analysis Shmuel Razin and David Yogev E5 Southern Blot Analysis and Ribotyping David Yogev and Shmuel Razin E6 Phylogenetic Classification of Plant Pathogenic Mycoplasma-like Organisms or Phytoplasmas Bernd Schneider, Frich Seemueller, Christine D. Smart, and Bruce C. Kirkpatrick E7 Determination of Cholesterol and Polyoxyethylene Sorbitan Growth Requirements of Mollicutes Joseph C. Tully
SECTION F
PATHOGENICITY F1 Introductory Remarks Shmuel Razin F2 Mycoplasma Adherence to Host Cells: Methods of Quantifying Adherence Itzhak Kahane and Fnno Jacobs F3 Mycoplasma Adherence to Host Cells: Epitope Mapping of Adhesins Fnno Jacobs
XIX
XX
Contents of Volume I F4 Oxidative Damage Induced by Mycoplasmas Itzhak Kahane F5 Activation of Macrophages and Monocytes by Mycoplasmas Ruth Gallily, Ann Avron, Gerlinde jahns-Streubel, and Peter F. Muhlradt F6 Identification, Characterization, and Purification of Mycoplasmal Superantigens Barry C. Cole and Curtis L Atkin F7 Mycoplasmal B-Cell Mitogens Yehudith Naot F8 Modulation of Expression of Major Histocompatibility Complex Molecules by Mycoplasmas P. Michael Stuart and Jerold G. Woodward F9 Interaction of Mycoplasmas with Natural Killer Cells Wayne C. Lai and Michael Bennett Index
MOLLICUTE-HOST INTERRELATIONSHIPS: CURRENT CONCEPTS AND DIAGNOSTIC IMPLICATIONS Joseph G. Tully
Introduction For almost four decades after their discovery, mollicutes (then called pleuropneumonia or pleuropneumonia-like organisms) were considered to be associated only with animal hosts as pathogens or commensals. The development of such a concept was reasonable when one examines the historical reports from the time of the cultivation of the first mollicute in 1898, the agent of contagious bovine pleuropneumonia, to the discovery of other mollicutes in other animal hosts (sheep, goats, rodents, dogs, and poultry) in the era between 1920 and 1937. As would have been expected, the report of a mollicute in the lower genital tract of a human female (Dienes and Edsall, 1937; Dienes, 1940) had a major impact on the ideas of host relationships of these organisms. However, the presence of competitive bacterial flora in humans and in many of the animal tissues being examined at the time was a major obstacle to further isolation and characterization of the mollicute flora of such hosts. The development of improved culture media, containing bacterial inhibitors for both gram-positive and gram-negative bacteria (Edward, 1947), considerably advanced the ability to cultivate and identify mollicutes from primary host tissues (Smith and Morton, 1951). This major technical accomplishment, coupled with the development of modified culture medium formulations and cultivation attempts on a variety of
Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
2
Joseph G. Tully
different hosts and tissues, including plants and invertebrates, has resulted in a continual expansion in the numbers of known mollicute species. At this writing, more than 158 mollicute species have been formally described, with possibly a dozen or more candidates in waiting. The known host range of mollicutes currently includes humans, nonhuman primates, a variety of domestic and wild animals and birds, and a large and expanding number of plant and insect species. The individual host relationships extend from a prominent role as important primary or opportunistic pathogens to that of commensals or epiphytes, and some mollicutes might also exist as saprophytes. This introductory chapter attempts to provide a concise review of the known distribution of mollicutes, in which some current concepts on various levels of mollicute-host interrelationships and application of this information to current approaches to laboratory diagnosis of mollicute infections are discussed. More extensive coverage and documentation of historical developments and the epidemiological data discussed here can be found in the five-volume series of The Mycoplasmas (Barile and Razin, 1979; Tully and Whitcomb, 1979; Whitcomb and Tully, 1979, 1989; Razin and Barile, 1985), two earlier mycoplasma methodology volumes (Razin and Tully, 1983; Tully and Razin, 1983), and selective reviews, symposia, or books (Tully, 1985; Razin, 1991; Tully and Whitcomb, 1991; Maniloff er a/., 1992; Kahane and Adoni, 1993; Cassell, 1993; Whitford et aL, 1994; Blanchard and Montagnier, 1994).
Acquisition and Transmission Mechanisms of Mollicutes As with many other biological entities, acquisition of mollicutes can be associated with primary transmission involving direct contact of host with host or transfer mediated through secondary means, such as aerosols or fomites, food and/or water, insect vectors or other carriers, and nosocomial acquisition (e.g., organ or tissue transplants). This discussion considers only broad areas of mollicute acquisition since more detailed information on the means by which mollicute species might be acquired and disseminated are covered later or in other sections of the volume. Many mollicutes are inhabitants of the mucous membranes of the respiratory, urogenital, or gastrointestinal tracts of vertebrates, so that direct host to host transmission of organisms in man and in many animals occurs through oral to oral, genital to genital, or oral to genital contact. Certain mollicutes that are part of the normal flora of the oropharynx and lower genital tract are most likely acquired by oral to oral or genital to genital contact. However, changes in sexual mores have resulted in apparent alterations in the host tissue location of mollicutes, so that some mollicutes commonly found in the oropharynx can occur in the urogenital tract and vice versa (See below). In some predatory insects, the
Mollicute-Host Interrelationships
3
organisms are acquired directly by eating other colonized or infected hosts. Direct host to host acquisition of mollicutes, particularly Ureaplasma urealyticum, has also been documented in neonates through in utero infection (see Chapter D4, this volume). Secondary transmissions of mollicutes are probably acquired most frequently through respiratory aerosols or fomites, as evidenced by the wide occurrence of mollicutes as etiologic agents of the many acute respiratory diseases of man and animals. Some of the normal commensal mollicute flora of the oropharynx of man and animals also may be acquired by this means. Food (and possibly water) obtained through feeding excursions plays a major role in the acquisition of mollicutes by insects, where the organisms are present in plant sap or nectar, or where such materials are contaminated with other insect-derived excretions. Food also plays a major role in dissemination, colonization, and transfer of nonpathogenic or infectious animal mollicutes from adult to the young through mammary (milk) transmission. In the transmission of mollicute diseases (citrus stubborn, com stunt, and phytoplasma infections) from plant to plant, insect vectors play a predominant role. These transmissions usually involve a clearly defined biological cycle within the insect, concerning acquisition, multiplication in various tissues (adipose tissue, salivary gland, etc.), and eventual reinoculation into susceptible plants. Secondary transmission of mollicutes can also occur through healthy carriers of the organisms or through convalescent carriers recovering from acute infections, as apparently occurs in some Mycoplasma pneumoniae respiratory infections in humans and probably in other acute and chronic respiratory diseases in domestic animals. Infected carriers also play an important role in the transmission of Spiroplasma melliferum and S. apis infections from foraging bees to their hives. Also, mollicute transmission after plant grafting of infected tissue to healthy rootstock, or through human organ or tissue transplantation, has been described. The presence of mollicutes in semen is important in sexually transmitted diseases in a variety of hosts and has become a critical complication in artificial insemination programs in humans and bovines (see Chapters D3, D6, and E7, this volume). Contamination of semen has had important consequences, including transmission of mollicutes and mollicute infections to other hosts, direct detrimental effects on successful host fertilization, and introduction of both pathogenic and nonpathogenic mollicutes to other geographic areas.
Host Colonization versus Host Infection with Mollicutes Since many mollicutes possess the ability to adhere to epithelial surfaces, any mechanism that promotes the direct or indirect exposure of the organism to susceptible tissue sites (e.g., respiratory, urogenital, and gastrointestinal tissues)
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Joseph G. Tully
readily results in colonization. Adherence mechanisms are well developed in many of the pathogenic mollicutes with organized attachment structures and with specific adhesin proteins on their exterior membrane surface (see the introductory chapter and Chapters F2 and F3 in Vol. I). However, other mollicutes, particularly those considered to be commensals in the oropharynx and urogenital tracts of humans and other animals, also possess mechanisms for cell adherence without well-defined tip structures or established adhesin proteins. The cytadherence mechanism for such mollicutes appears to involve the presence of specific glycolipid and/or glycoprotein receptors on the eukaryotic host cell surface (Razin and Jacobs, 1992). Early studies have shown that the human upper respiratory and urogenital tracts of human newborns are first colonized with mollicutes acquired during passage through the female birth canal. However, as noted earlier, there is now substantial evidence that, on a rare occasion, mollicute colonization can occur in utero, again possibly related to transmission or acquisition of mollicutes (M. hominis and U. urealyticum) in the lower female urogenital tract at the time of fertilization or during early pregnancy. In the usual situation, neonatal colonization with both human oral and genital mollicutes generally does not persist, and the number and type of mollicutes decrease with age. However, welldocumented studies have shown that subsequent reacquisition of a urogenital mollicute flora in humans is directly related to sexual contacts. Likewise, it is assumed that oropharyngeal commensals are reacquired later through oral to oral transmission. Less information is available on the mechanisms and timing of mollicute colonization of newborn vertebrates, including birds, and insects or other arthropods. In vertebrates, acquisition from the maternal genital tract, and in utero transmission (including egg transmission in the case of birds), apparently represents the initial mechanism. Colonization of arthropod and insect hosts probably occurs most frequently during feeding excursions on plant surfaces, flowers, and on nectar (Hackett and Clark, 1989). However, transovarian transmission of spiroplasmas has been well established in Drosophila species (Williamson and Poulson, 1979). The normal persistent mollicute colonization of major anatomical sites in humans and numerous animal hosts has created major complications in efforts to define a relationship between organism and disease. To avoid such complications in experimental studies, investigators have utilized gnotobiotic or specific pathogen-free (SPF) hosts that are defined as free of mollicutes (see chapters in Section E, this volume). However, while experimental infections in a variety of large and small animal hosts have helped to establish the pathogenicity of many host-restricted mollicutes for the respiratory tract, difficulties have not been overcome in producing and maintaining large gnotobiotic or SPF animals free of urogenital tract mollicutes.
Mollicute-Host Interrelationships
5
As with many other microbe-host interrelationships, mollicutes can be found on the mucosal surface as part of the normal flora or the host may acquire a pathogenic species able to colonize, invade host tissue, and eventually produce tissue damage. Some of the mechanisms that are thought to be involved in mollicute pathogenicity are outlined in the introductory chapter in Vol. I. Host factors also play an important and complex role in determining whether an acquired mollicute behaves as a pathogen or is avirulent. Although most of these factors are not well defined or understood at this time, it is clear that endogenous mollicutes can become pathogenic when the human host immune system has been compromised or when surgical intervention or trauma allows normal mollicute commensals to enter the circulation or other host tissues. In these occurrences, such common commensals as M. hominis, M. salivarium, and U. urealyticum (Furr et al., 1994; Gelfand, 1993; Meyer and Clough, 1993) have been shown to induce septicemia and overt mollicute invasion of various host tissues and organs, with joint localization being especially prominent.
Host Specificity of Mollicutes One of the hallmarks of the mollicute-host relationship recognized by early workers was the remarkable host specificity of the organisms. The earliest isolates found in bovine pleuropneumonia had not at the time been isolated from goats or sheep, rodents, or from other animal or human hosts examined shortly thereafter. Although this striking host specificity has for the most part remained a characteristic feature of mollicutes, exceptions have been discovered as new mollicutes were isolated and the known host range expanded. Strains of M. arginini, M. canis, and many established Acholeplasma species have been found to occur in a wide variety of different animal hosts. Only a few examples of some recent unusual host relationships will be noted here since the earlier references and textbooks mentioned can provide additional background. Several other reports further support the apparent transmission of animal mollicutes to humans. The few earlier reports of the possible transmission of M. arthritidis and M. canis to humans have been reviewed (Armstrong et al, 1971; Tully, 1993). Of more immediate concern are instances where serious clinical infections in humans have occurred, including those with a fatal outcome, from apparently animal-derived mollicutes. McCabe and associates (1987) described a mycoplasmal infection in the hand of a veterinarian acquired through a cat bite. The infection resulted in severe soft tissue cellulitis with tissue destruction sufficient to require a tendon graft. The organism (strain M7806) was identified at the time as an unclassified, glucose-fermenting Mycoplasma species. Although other serologically related strains were later isolated from the oropharynx of felines, the
6
Joseph G. Tully
organism remains unclassified. Yechouron and colleagues (1992) presented observations on a fatal human infection with M. arginini. This organism was repeatedly isolated from the blood and bronchial washings of the patient during the course of the infection. The fact that the patient had advanced Hodgkin's disease and a marked immunodeficiency obviously contributed to the invasion of the organism and to the inability to control the infection with appropriate chemotherapy. Although the source of the infection was not determined through epidemiologic or occupational studies, the most likely acquisition was thought to have involved aerosol transmission during patient employment in a large animal slaughterhouse. Examples of the possible reverse transfer of mollicutes of human origin to animal hosts, and the somewhat less unusual cross transfer of mollicutes among certain animal hosts, form another part of the picture of changes in host specificity of mollicutes. M. salivarium, a species usually confined to humans or other primates, was repeatedly isolated from nasal and pharyngeal secretions of swine (Erickson et al., 1988). Mycoplasmas usually found in avian and bovine hosts have been reported to occur in swine (Taylor-Robinson and Dinter, 1968), and two mollicutes primarily of rodent origin, M. arthritidis andM. collis, have been isolated from synovial fluids of arthritic swine (Binder et al., 1990). Furthermore, the demonstration of a large number of isolates of a canine mycoplasma from the respiratory tract of calves also appears to indicate some changes in the usual host relationships. More than 40 mycoplasma strains were isolated from the lungs of calves in 19 herds in The Netherlands (ter Laak et al., 1993). Initially, these strains were thought to be new species since extensive serologic testing did not indicate that they were related to any established mollicute. However, subsequent investigations with other serologic techniques (agar plate immunofluorescence tests) and a comparison of restriction endonuclease patterns to mollicutes with identical DNA base composition (G + C) values indicated the likelihood that these strains were closely related to M. canis. The failure of initial tests to detect the serologic relationship was thought to be due to the extensive serologic heterogeneity among M. canis strains, first described more than 40 years earlier. Retrospectively, it was also determined that M. canis strains had been identified in the respiratory tract of calves in Canada as early as 1974 (H. L. Ruhnke and J. G. Tully, unpublished studies). The presence of mollicutes in plants and insects provides an opportunity for their possible transfer to human or other animal hosts and for the reverse transmission to occur. The isolation and identification of an avian mollicute (M. iowae, strain PPAV) from apple seeds has become not only an example of a mollicute in association with an unusual host transfer, but a classic demonstration of the application of 16S rDNA gene sequence analysis in mollicute identification. Details of the isolation and earlier difficulties in identification of the organism, including an extensive serological analysis, have been described (Grau et al, 1991; Tully, 1989). Again, the initial failure to detect the apparent
Mollicute-Host Interrelationships
7
serologic relationship was thought to relate to serologic heterogeneity among strains of M. iowae. In efforts to explore possible genetic relationships of PPAV to other mollicutes, the full 16S rDNA was sequenced. A search of gene data banks showed a difference of only five bases in the total 16S rDNA of PPAV and the type strain (695) of M. iowae. DNA/DNA hybridization, genome size comparisons, and other molecular and phenotypic comparisons later confirmed the identification of PPAV as M. iowae. An explanation of how this avian moUicute apparently occurred in seeds removed from an apple under aseptic isolation techniques is not readily apparent. Another example of possible transmission from animal host to plant was uncovered in the case of the plant-derived F5 strain (J. C. Vignault, C. Saillard, P. Carle, J. M. Bove, D. L. Rose, and J. G. Tully, unpubUshed studies; Tully, 1989). This nonhelical, sterol-requiring organism was isolated from a fieldcollected wild lettuce plant near Bordeaux, France in 1981. It was characterized extensively in comparative studies with other plant/insect mollicutes and appeared to be distinct from all other mollicutes in that habitat. However, tests to define an optimal growth temperature indicated the strain grew best at 37°C. Subsequent reciprocal serologic tests (growth inhibition and epi-immunofluorescence) with antisera to then current Mycoplasma species indicated that strain F5 was closely related to the T37 strain of M. equigenitalium. A comparison of genomic properties and biochemical activities of strains F5 and T37 (G + C of 31.0 mol%, genome of 880 kbp, positive glucose fermentation, negative arginine hydrolysis, positive film and spot reaction) provided additional confirmation of this identification. Since M. equigenitalium is a normal inhabitant of the equine genital tract, it was speculated that the most likely source for plant contamination by this mollicute was equine urine or other urogenital-derived material.
Habitat and Ecology of Mollicutes Distribution in Humans and Nonhuman Primates
Mycoplasmas, ureaplasmas, and a few acholeplasmas have been isolated from the oropharynx and lower urogenital tract of most humans (Table I) (Tully, 1993). The first five species listed in the table are considered to be representative of the normal human oropharyngeal flora. M. salivarium appears to be the most frequent mollicute in the oral cavity, with incidences of 60-80% in various adult populations, whereas M. orale can usually be found in 30-60% of adult throats. The other human oropharyngeal mollicutes are isolated only infrequently. The occurrence of M. salivarium in the human genital tract has been reported twice, and more than 20 isolates of M. pneumoniae were cultivated from cervical
Joseph G. Tully
TABLE I PRIMARY AND SECONDARY HUMAN TISSUE SITES OR ORGANS COLONIZED OR INFECTED WITH MOLLICUTES"
Species
Primary site
Mycoplasma salivarium M. orale
Oropharynx Oropharynx
M. buccale M. faucium M. lipophilum M. pneumoniae
Oropharynx Oropharynx Oropharynx Oropharynx; lung; pleural fluid; bronchoalveolar lavage fluid
M. genitalium
Urogenital tract; oropharynx Lower genital tract; oropharynx
M. hominis
M. fermentans
Lower genital tract; respiratory tract
M. primatum M. spermatophilum M. pirum
Female urethra Cervix/sperm Peripheral blood cells? ?
M. penetrans Ureaplasma urealyticum
Acholeplasma laidlawii A. oculi
Genital tract; oropharynx; bronchoalveolar lavage fluid; placenta 7 7
Secondary site Cervix/vagina; arthritic joint Leukemic bone marrow; lymph node and skin (in sarcoidosis)
Arthritic joints; skin lesions (in StevensJohnson syndrome; middle ear fluid; cerebrospinal fluid; tuboovarian abscess; cervix and vagina; pericardial fluid; heart blood, kidney, and brain at autopsy Synovial fluid in arthritic joint Lung and pleural effusion; blood in postpartum septicemia; prostheses infections; organ and tissue transplant infections; trauma/surgical site infections; malignancies; cerebrospinal fluid; peritoneum; synovial fluid (arthritis); skin; pericardium; amniotic fluid; neonatal septicemia Leukemic bone marrow; arthritic joints; peripheral blood lymphocytes in acquired immunodeficiency syndrome (AIDS); urine in AIDS Umbilicus Peripheral blood lymphocytes Urine (male) in AIDS Lung and lower respiratory tract (in pneumonia); blood in neonatal septicemia; cerebrospinal fluid; transplants; surgical sites, arthritic sites; renal calculi; amniotic fluid; postpartum septicemia; neonatal lung and brain Oropharynx; bum infections; vagina Amniotic fluid
"Modified from Tully, J. G. (1993). Current status of the mollicute flora of humans. Clin. Infect. Dis. 17 (Suppl. 1), S2-S9, by permission of the University of Chicago. Copyright © 1993 The University of Chicago Press.
Mollicute-Host Interrelationships
9
specimens of women attending various gynecological clinics in Canada (Goulet et al., 1995). The implications of these reports is that transmission had occurred through oral-genital contact. M. hominis and U. urealyticum are the most frequently encountered mollicutes in the lower urogenital tract of humans, and their recurrence and incidence after puberty are correlated with increasing sexual contact, especially to the number of sexual partners (McCormack et al, 1973). Rates of M. hominis colonization are lower in men than in women; in a survey of women attending a sexual disease clinic, a colonization rate as high as 94% was recorded (TuUy et al., 1983). The clinical roles that M. hominis and U. urealyticum play in human opportunistic infections, urogenital disease, and neonatal infections, and their laboratory detection, are discussed in Chapters D3 and D4 of this volume. The other major pathogens in the group, or those currently considered to have an association with human disease, include M. pneumoniae, M. fermentans, M. genitalium, and M. penetrans (for a discussion of individual species, see Chapters D2 through D5, this volume). M. pirum, which was first found as an in vitro cell culture contaminant (see chapters in Section F, this volume), has been isolated from human peripheral blood cells and is now tentatively considered to be of human origin (Blanchard and Montagnier, 1994) (see also Chapter D5, this volume). Although few isolates of M. primatum or M. spermatophilum have been reported, they appear to be part of the human urogenital flora. The few acholeplasmas found in humans and nonhuman primates may represent the occasional transmission from other animal hosts since some acholeplasmas have been isolated regularly from domestic animals and birds and others from plants and insects (see Waites et al., 1987 and later discussion). The mollicute flora of a considerable number of nonhuman primates has been explored, primarily from the standpoint of their use as experimental models for pathogenicity studies on microbial agents, including mycoplasmas of human origin. An extensive listing of mollicute species identified within various nonhuman primates (Somerson and Cole, 1979) included M. hominis, M. fermentans, M. salivarium, M. orale, M. buccale, M. faucium, M. primatum, M. lipophilum, M. moatsii, M. canis, M. arthritidis, Acholeplasma laidlawii, and some ureaplasmas. Most of the isolates were from the oropharynx, but M. moatsii and some M. salivarium strains occurred in the urogenital tract (see also Hill, 1983). A new species described as M. indiense has been isolated from the throats of a rhesus monkey and a baboon (Hill, 1993). Distribution in Large Domesticated Animals BOVINES
Twenty-three species of mollicutes occur regularly in cattle, including 14 Mycoplasma, 3 Acholeplasma, 1 Ureaplasma, and 5 AnaeroplasmalAsteroleplasma species (Gourlay and Howard, 1983; Robinson and Freundt, 1987; see
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Joseph G. Tully
also Chapter D6, this volume). Most isolates are found in the upper respiratory tract, ocular sites, or the genital tract. Species isolated from mammary sites also are frequently found in respiratory or urogenital tracts. The obligate anaerobic anaeroplasmas and asteroleplasmas have been found only in the bovine rumen (Robinson and Freundt, 1987). The list of species found in bovines has changed only slightly since an earlier review (Gourlay and Howard, 1983) and, as noted at that time, several equine, feline, or avian mollicutes (M. equirhinis, M. galeae, M. gallisepticum, and M. gallinarum) also have been recovered from bovines (see also Lauerman, 1994; Jasper, 1994; Ruhnke, 1994). EQUINES
The equine mollicute flora is generally considered to consist of 11 species from urogenital or respiratory tracts. (M. equigenitalium, M. equirhinis, M. fastidiosum, M.felis, M. subdolum, A. equifetale, A. hippikon, A. laidlawii, A. modicum, A. multilocale, A. oculi and A. parvum), with occasional isolates of M. arginini, M. feliminutum, M. pulmonis, M. salivarium, A. axanthum, and A. granularum also being made from these sites in horses (Hill et aL, 1992; Whitford and Lingsweiler, 1994). CAPRINES/OVINES
Twenty mollicute species occur regularly in goats and sheep, including 16 Mycoplasma and 4 Acholeplasma species, with another 2 unclassified groups (strain 2D and related mycoplasmas, and a ureaplasma cluster) representing putative species. A list of species, including biologic characteristics and isolation sites, is given in Chapter D7 of this volume (see also Rosendal, 1994). Species of the obligately anaerobic genera Anaeroplasma or Asteroleplasma also occur in the rumen of sheep (Robinson and Freundt, 1987). PORCINES
The principal mollicute flora of swine include M. hyorhinis, M. hyopneumoniae, M. hyopharyngis, M. hyosynoviae, and M. flocculare, with occasional isolations of M. arginini, M. bovigenitalium, M. buccale, M. gallinarum, M. iners, M. mycoides subsp. mycoides, M. salivarium, M. sualvi, A. axanthum, A. granularum, A. laidlawii, and A. oculi, and some unclassified ureaplasmas or obligately anaerobic mollicutes (see Chapter D8, this volume; Ross and Whittlestone, 1983; Binder and Kirchhoff, 1988; Armstrong, 1994). Distribution in Small Domesticated Animals and Birds CANINES/FELINES
The canine mollicute flora consists of at least 14 species, including primarily M. canis, M. cynos, M. edwardii, M. maculosum, M. molare, M. opalescens.
Mollicute-Host Interrelationships
11
M. spumans, and U. canigenitalium, with occasional isolates of M. arginini, M. bovigenitalium, M. felis, M. feliminutum, M. galeae, A. laidlawii, an unclassified isolate (HRC 689), and several serologically distinct groups of ureaplasmas (Ogata, 1983; Whitford and Lingsweiler, 1994). Smaller numbers of mollicute species have been isolated from cats, with the principal flora consisting of M. felis, M. feliminutum, M. galeae, Ureaplasmafelinum, and U. call, and a currently unclassified group of mycoplasmas (M7806) (H. L. Ruhnke, personal communication). Other reported isolations include M. arginini, M. arlhrilidis, M. galliseplicum, and M. pulmonis (Ogata, 1983). SMALL LABORATORY ANIMALS
Mice and rats generally share a characteristic mollicute flora, consisting of M. arlhrilidis, M. collis, and M. pulmonis. Although mice also have both M. neurolylicum and M. muris as part of their normal flora, these two mollicutes so far have not been identified in rats (Cassell el ai, 1983; Davidson el al., 1994). Most of the mollicutes are maintained in the respiratory or urogenital tracts, but many also can be isolated from conjunctivae, middle ear, and brain. Guinea pigs harbor strains of M. caviae, M. cavipharyngis, A. cavigenilalium, A. granularum, and A. laidlawii (Hill, 1983), with most of the species occurring in the urogenital tract. Chinese hamsters are reported to have both M. criceluli and M. oxoniensis in the conjuctivae (Hill, 1991). Few mollicutes have been identified in rabbits, although earlier isolation of M. pulmonis was reported from rabbits housed near rodent colonies, and a new species (A. mullilocale) has been isolated from rabbit feces (Hill, 1992). AVIAN
The widespread occurrence and distribution of mollicutes in avian hosts is reviewed in Chapter D9 (this volume). Tv^tniy-i^o Mycoplasma species and U. gallorale have been identified in various birds, with many of the more recent isolates from birds of prey. Additional information on tissue locations of these mollicutes can be found in other reviews (Jordan, 1983; Kleven, 1994). Distribution in Wild Animals and Aquatic Hosts The occurrence of mollicutes in many types of domestic animals prompted questions about the possible distribution of the organisms in wild animals or aquatic hosts. Thus, a considerable mollicute flora has now been defined in a variety of wild animals, particularly those housed in zoos, as well as in several aquatic hosts. Some references to earlier isolations can be found in reports of the International Research Programme on Comparative Mycoplasmology (1990) or in a review (Razin, 1992), including M. cilelli from the trachea, lung, andhver of ground squirrels (Rose el al., 1978); M. mobile from a fresh water trench, and
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Joseph G. Tully
M. phocarhinis and M. phocacerebrale from the upper respiratory tract and tissues of harbor seals (Giebel et al., 1991). Another harbor seal isolate from lung and trachea also was described as M. phocidae (Ruhnke and Madoff, 1992). M. testudinis was isolated from the cloaca of a pet turtle (Hill, 1985) and M. mustelae from the lungs of mink (Salih et al, 1983). Previously unknown mollicute species have been found in many wild felines, including M. felifaucium from the respiratory tract of a puma (Hill, 1986), M. leopharyngis and M. simbae from the lion pharynx, and M. leocaptivus from the throats of both a lion and a leopard (Hill, 1992). Two additional moUicutes, apparently distinct from other established species, are currently being described as new species: a urogenital isolate from an elephant (H. Kirchhoff, personal communication) and several related strains isolated from the desert tortoise (M. Brown, personal communication). Distribution in Arthropods^ Including Insects Since the initial discovery of moUicutes in insect-transmitted phytoplasma plant diseases in 1967 and the isolation and characterization of the first helical mollicute {Spiroplasma citri) in citrus stubborn disease, insects and other arthropods have been a rich source of moUicutes. The organisms include numerous species and group representatives now assigned to the genus Spiroplasma and to an expanding number of species in the newly established genera of Entomoplasma and Mesoplasma (Tully et al., 1993). Since the information on this topic is also extensive, key references to work prior to 1991 can be found in earlier books and reviews (Whitcomb and Tully, 1979; Razin and Tully, 1983; Tully and Razin, 1983; Tully et al, 1987; Tully, 1989; Whitcomb and Tully, 1989; Williamson et al, 1989; Hackett et al, 1990; Tully et al, 1990; Tully and Whitcomb, 1991). The Appendix in this volume contains a list of currently recognized species and type strains of arthropod-associated moUicutes. In the case of spiroplasmas, the list includes group representatives that have not yet received species names. Spiroplasmas are found most frequently in the insect gut, less frequently in hemolymph, and occasionally in salivary glands. The organisms are generally acquired by natural feeding, from plant tissue, nectar, or insect excretions, or by ingestion of other insects. As noted earlier, at least one example of transovarial transmission of spiroplasmas (in Drosophila) is known. Most spiroplasmas are found in six evolutionarily advanced insect orders: Hymenoptera, bees and wasps; Coleoptera, plant- and flower-feeding beetles, firefly beetles, etc.; Diptera, flower-feeding flies, tabanids (horseflies), mosquitoes, fruit flies, etc.; Lepidoptera, butterflies; Homoptera, leafhoppers; and Hemiptera, green-leaf bugs. A serologically distinct spiroplasma (PALS-1) has been identified in the gut of a dragonfly, at this time the single representative from a primitive insect order (Odonata) found to harbor spiroplasmas. Two species {Spiroplasma mirum
Mollicute-Host Interrelationships
13
and S. ixodetis) have been isolated from tick genera (Haemaphysalts and Ixodes, respectively), but tissue localization within ticks and conditions involved in acquisition and maintenance are not well known. Some insect hosts of spiroplasmas function as important vectors in the transmission of plant or insect diseases. Leafhoppers infected with S. citri are the principal vectors for the transmission of citrus stubborn disease, not only among citrus plants but also among other infected plants or weeds and citrus. Likewise, leafhoppers infected with S. kunkelii are the principal vectors for the transmission of com stunt disease. Honeybees infected with either S. melliferum or S. apis are the major carriers and vectors for the transmission of lethal spiroplasma diseases to other honeybees or to their hives. Techniques for diagnosis of spiroplasmal infections in insects and experimental infections in such hosts are presented in Chapters DIO, D12, and E8 (this volume). Most nonhelical mollicutes of the genera Entomoplasma or Mesoplasma have been found as inhabitants of the insect gut. This situation may reflect an emphasis on the ecology of insect mollicutes. However, some entomoplasmas and mesoplasmas have now been identified also on plant surfaces (see later), and it is expected that most species in either group are acquired from various plant surfaces or flowers through insect visitation and feeding excursions. E. ellychniae was found in the hemolymph of a firefly, whereas E. somnilux, E. lucivorax, and E. luminosum appear to be gut inhabitants of other firefly species (Williamson et al., 1990; Tully et al., 1993). In contrast, E. melaleucae was first isolated from several species of subtropical flowers in 1979 and was only later found in a bee. The current distribution of Mesoplasma species, which were initially described as acholeplasmas (Tully et al., 1993), has been summarized (Tully et al., 1994b). Again, most isolates have been made from gut fluids of insect hosts, with a few isolations also from plant surfaces. The first species described in the group {Me. florum) was found on flower surfaces, but subsequent strains have been identified in the gut and hemolymph of beetles. Me. entomophilum is widely distributed among insects, occurring as part of the gut flora of more than nine different insect genera. The only occurrence of Me. seijfertii strains in insects has been reported in isolations made from the gut contents of Aedes mosquitoes and a Chrysops fly in France (Chastel et al., 1994). With the exception of Me. lactucae, the balance of the currently recognized Mesoplasma species (see Table IV in the Appendix) are inhabitants of a variety of insects (fireflies, horseflies, other beetles, bees, butterflies, etc.). The initial reports describing non-sterol-requiring mollicutes in insects as Acholeplasma species were eventually corrected when it was established that these mollicutes (now termed mesoplasmas) were phylogenetically distinct from classic acholeplasmas (Tully et al., 1993). Currently, conventional acholeplasmas have not been identified as part of the insect or arthropod flora. However, since at least four or five Acholeplasma species have been found on a
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Joseph G. Tully
variety of plant surfaces (see later), it is more than likely that these mollicutes also are part of the insect flora. Finally, phloem-feeding insects (mainly leafhoppers) are the primary hosts and vectors of a large collection of plant pathogenic mollicutes, including a few spiroplasmas and many phytoplasmas (see McCoy et aL, 1989; Chapter E6, in Vol. I; and Chapter E9, this volume).
Distribution in Plants
The largest group of mollicutes inhabiting plants is the currently uncultivable cluster of plant-pathogenic organisms now called phytoplasmas. These mollicutes are transmitted primarily by phloem-feeding leafhoppers and are the causative agents of more than 300 different plant "yellows" diseases. Genetic and taxonomic studies have indicated that these organisms have a close evolutionary relationship to the acholeplasmas and that DNA from more than 51 representative phytoplasmas can be placed into 12 distinct groups (see Chapter E6, Vol. I; Chapters DU and E9, this volume; and McCoy et al, 1989). Spiroplasma occurrence in plant hosts is based either on invasion of the plant sieve tubes in the course of a biological cycle involving the feeding activities of homopterous insects (leafhoppers) or as external contamination of floral parts from deposition by flower-visiting insects. Only spiroplasma invasion into plant sieve tubes results in disease, as is seen in various citrus plant diseases with S. citri, in com stunt disease with S. kunkelii, and in the so-called periwinkle disease due to infections with S. phoeniceum (see Chapters DIO and E8, this volume). Numerous other Spiroplasma species (or groups) have been isolated from flower surfaces and nectar, but no evidence is currently available that these organisms also occur within internal plant tissues (see Table V in Appendix). However, some normal flower-inhabitating spiroplasmas acquired by specific insects function as pathogens, as in the case of S. apis strains acquired by honeybees. The only isolations of Entomoplasma species from plants so far have come from tree flowers; these include flowers of two species of the genus Melaleuca and the silk oak (Grevillea sp.) in Florida. Tropical flowers (grapefruit, lemon, and powder puff plant) have also been the source of three strains of Me. florum. This organism and Me. entomophilum occur frequently in a variety of insects, which explains their repeated isolation from various wild plants (aster, sunflower, gumweed, goldenrod, etc.) in Colorado and Nebraska. The remaining mesoplasmas have not been found often in plant hosts, which also appears to be reflected in a limited number of isolations of these organisms from insects. The reported isolations include Me. seiffertii from flowers of a sweet orange tree in Morocco and wild angelica in France, Me. lactucae from lettuce, and Me. coleopterae from flowers of the vera dulca tree in Texas.
Mollicute-Host Interrelationships
15
The occurrence of classic acholeplasmas on plant surfaces is somewhat enigmatic since none of the acholeplasmas identified so far has been isolated from insects. A. axanthum has been identified on vegetable plants, on wild plants (mesquite, Prunus sp., creosote bush) in the southwestern United States, and on crown tissues of coconut palms with lethal yellowing disease (phytoplasma) in Jamaica. Both A. oculi and A. palmae have been isolated from similar coconut palm crown tissues in Jamaica, and A. laidlawii and A. brassicae occur as surface contaminants on vegetables (Tully et al., 1994a).
Habitat Consideration and Diagnostic Laboratory Approaches One of the most interesting undercurrents running through the foregoing discussion is the possibility that mollicutes might be able to cross species barriers much more frequently than had been expected from earlier experiences. Whether such breaks represent actual changes in host relationships or the recognition of a few mollicutes that might be less host-specific than others is still uncertain. However, experiences with unusual host transfers suggest that it would now seem prudent to develop broader approaches for laboratory diagnostic techniques for mollicutes that would emphasize rapid differentiation to the genus level. After a preliminary screening and separation, other conventional techniques (serological, etc.) could then be employed for a more critical differentiation to the species level. A forward step in this direction has been outlined in a discussion of diagnostic procedures for arthropod mollicutes (Chapter D12, this volume). This approach is based on the description of new taxonomic groups of mollicutes identified in plant and insect hosts (Tully et al., 1993; see also Chapter E2, Volume I). The following discussion expands this idea to mollicutes of human and animal origin and details practical techniques required for final identification. Since the obligate anaerobic mollicutes (members of the genus Anaeroplasma and genus Asteroleplasma) require specialized equipment for isolation and growth, and the phytoplasmas are presently uncultivable, these organisms are not considered here. A revision of recommended techniques for describing new mollicutes will provide additional references for methods discussed next (International Committee on Systematic Bacteriology, 1995). A schematic approach to the differentiation of mollicute genera is presented in Table II. A preliminary requirement is that the organism to be identified can be cultivated in some type of broth and agar media. Growth in liquid medium will vary from heavy to light, but large amounts of sediment should raise suspicions of a bacterial agent. Several mollicute species require anaerobic conditions for growth in liquid medium, so initial attempts at cultivation should include this environment. Classic fried egg-type agar colonies are frequently not observed
16
Joseph G. Tully TABLE II SCHEMATIC APPROACH TO LABORATORY DIFFERENTIATION OF MAJOR GENERA OF CLASS MOLLICUTES
A. Preliminary characteristics to be determined Growth on hquid and solid medium (preliminary biochemical properties) Purification by filtration cloning (3 X) (450-nm membrane filters) Filterable through 450-, 300-, 220-, and 100-nm membrane filters Growth in presence of penicillin and failure to revert to bacterial-type growth when grown in absence of antibiotics B. Morphology (dark-field microscopy or negative stain with electron microscopy) Helical > Genus Spiroplasma Nonhelical > Other moUicute C. Growth in serum-free medium < ^ > Genus Acholeplasma Positive Negative ^ Other moUicute D. Growth in serum-free medium containing 0.04% Tween 80 « —" Positive > Genus Mesoplasma Negative —» Other moUicute E. Optimal growth temperature <— ' ~ 30°-32°C > Genus Entomoplasma 35°-37°C > Genus Mycoplasma/Genus Ureaplasma F. Hydrolysis of Urea < > Genus Mycoplasma Negative > Genus Ureaplasma Positive
with many mollicutes (especially spiroplasmas), but most species are facultative anaerobes and will form colonies under anaerobic conditions. The organism should be filter-cloned at least three times to provide some assurance that the culture is not a mixture of more than one type of moUicute (see Chapter E2, Volume I). One of the most important preliminary features that should be observed in the characterization process is the ability of the organism to pass membrane filters of different porosities. The number of organisms in a broth culture at an early logarithmic phase of growth should be determined, usually by serial 10-fold dilutions for color-changing units per milliliter or by colony counts on agar. Filtrates of this same broth culture obtained after passage through 450-, 300-, 220-, and 100-nm membranes should be quantitated in a similar manner. At least some cells of a true moUicute should pass through 220nm filters, although the numbers may be considerably reduced from those in the unfiltered broth. These features, coupled with the ability of the organism to grow in the presence of penicillin and the lack of reversion to bacterial-type colonies on solid medium devoid of penicillin or other antibiotic agents, are necessary prerequisites to identification. Morphological differentiation of spiroplasmas is most easily determined by
Mollicute-Host Interrelationships
17
conventional dark-field microscopic techniques. Examination should be performed on broth growth in the early logarithmic phase and at various stages of growth since some Spiroplasma species may be nonhelical at various stages (usually in the stationary part) of the growth cycle. Details for establishing growth requirements for sterol or for testing the ability of strains to grow in various serum-free medium formulations are given in Chapter E7 (Volume I). Determination of optimum growth temperature differentiates Mycoplasma and Ureaplasma species of vertebrates, those growing best between 35° and 37°C, from Entomoplasma species of plants and insects with a growth optimum around 30°C (see Chapter E2, Volume I). Finally, tests for urea hydrolysis will distinguish the Ureaplasma from Mycoplasma species.
References Armstrong, C. H. (1994). Porcine mycoplasmas. In "Mycoplasmosis in Animals: Laboratory Diagnosis" (H. W. Whitford, R. F. Rosenbusch, and L. H. Lauerman, eds.), pp. 68-83. Iowa State Univ. Press, Ames. Armstrong, D., Yu, B. H., Yagoda, A., ajpd Kagnoff, M. F. (1971). Colonization of human by Mycoplasma canis. J. Infect. Dis. 124, 607-609. Barile, M. F., and Razin, S., eds. (1979). "The Mycoplasmas," Vol. 1. Academic Press, New York. Binder, A., and Kirchhoff, H. (1988). Isolation of anaerobic moUicutes from the intestine of swine. Vet. Microbiol. 17, 151-158. Binder, A., AumuUer, R., Likitdecharote, B., and Kirchhoff, H. (1990). Isolation oiMycoplasma arthritidis from the joint fluid of boars. J. Vet. Med. Ser. B 31, 611-614. Blanchard, A., and Montagnier, L. (1994). AIDS-associated mycoplasmas. Anna. Rev. Microbiol. 48, 687-712. Cassell, G. H., ed. (1993). The changing role of mycoplasmas in respiratory disease and AIDS. Clin. Infect. Dis. 17 (Suppl.l), SI-S315. Cassell, G. H., Davidson, M. K., Davis, J. K., and Lindsey, J. R. (1983). Recovery and identification of murine mycoplasmas. In "Methods in Mycoplasmology" (J. G. TuUy and S. Razin, eds.), Vol. 2, pp. 129-142. Academic Press, New York. Chastel, C , Saillard, C , Le Goff, F., Gros, O., Helias, C , Poutiers, F., and Bove, J. M. (1994). Serological and molecular characterization of Mesoplasma seiffertii strains isolated from haematophagous dipterans. lOM Lett. 3, 455. Davidson, M. K., Davis, J. K., Gambill, G. P., Cassell, G. H., and Lindsey, J. R. (1994). Mycoplasmas of laboratory rodents. In "Mycoplasmosis in Animals: Laboratory Diagnosis" (H. W. Whitford, R. F. Rosenbusch, and L. H. Lauerman, eds.), pp. 97-133. Iowa State Univ. Press, Ames. Dienes, L. (1940). Cultivation of pleuropneumonia-like organisms from female genital organs. Proc. Soc. Exp. Biol. Med. 44, 468-469. Dienes L., and Edsall, G. (1937). Observations on the L-organism of Klieneberger. Proc. Soc. Exp. Biol. Med. 36, 740-744. Edward, D.G. ff. (1947). A selective medium for pleuropneumonia-like organisms. J. Gen. Microbiol. 1, 238-243.
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Erickson, B. Z., Ross, R. F., and Bove, J. M. (1988). Isolation of Mycoplasma salivarium from swine. Vet. Microbiol 16, 385-390. Furr, P. M., Taylor-Robinson, D., and Webster, A. D. B. (1994). Mycoplasmas and ureaplasmas in patients with hypogammaglobulinaemia and their role in arthritis: Microbiological observations over twenty years. Ann. Rheum. Dis. 53, 183-187. Gelfand, E. W. (1993). Unique susceptibility of patients with antibody deficiency to mycoplasma infection. Clin. Infect. Dis. 17(Suppl.l), S250-S253. Giebel, J., Meier, J., Binder, A., Flossdorf, J., Poveda, J. B., Schmidt, R., and Kirchhoff, H. (1991). Mycoplasma phocarhinis sp. no v.. Mycoplasma phocacerebrale sp. nov., two new species from harbor seals {Phoca vitulina L). Int. J. Syst. Bacteriol. 41, 39-44. Goulet, A. M., Dular, R., Tully, J. G., Billowes, G., and Kasatiya, S. (1995). Isolations of Mycoplasma pneumoniae from the human urogenital tract. J. Clin. Microbiol, (in press). Gourlay, R. N., and Howard, C. J. (1983). Recovery and identification of bovine mycoplasmas. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 81-89. Academic Press, New York. Grau, O., Laigret, F., Carle, P., Tully, J. G., Rose, D. L., and Bove, J. M. (1991). Identification of a plant-derived mollicute as a strain of an avian pathogen, Mycoplasma iowae, and its implications for mollicute taxonomy. Int. J. Syst. Bacteriol. 41, 473-478. Hackett, K. J., and Clark, T. B. (1989). Spiroplasma ecology. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.). Vol. 5, pp. 113-200. Academic Press, San Diego, CA. Hackett, K. J., Whitcomb, R. F., Henegar, R. B., Wagner, A. G., Clark, E. A., Tully, J. G., Green, F., McKay, W. H., Santini, P., Rose, D. L., Anderson, J. J., and Lynn, D. E. (1990). Mollicute diversity in arthropod hosts. Zentralbl. Bakteriol. Hyg., Suppl. 20, 441-454. Hill, A. C. (1983). Recovery and identification of mycoplasmas from other laboratory animals (including primates). In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 143-147. Academic Press, New York. Hill, A. C. (1985). Mycoplasma testudinus, a new species isolated from tortoise. Int. J. Syst. Bacteriol. 35, 480-492. Hill, A. C. (1986). Mycoplasma felifaucium, a new species isolated from the respiratory tract of pumas. J. Gen. Microbiol. 132, 1923-1928. Hill, A. C. (1991). Mycoplasma oxoniensis, a new species isolated from Chinese hamster conjunctivas. Int. J. Syst. Bacteriol. 41, 21-25. Hill, A. C. (1992). Mycoplasma simbae. Mycoplasma leopharyngis, and Mycoplasma leocaptivus sp.nov., isolated from lions. Int. J. Syst. Bacteriol. 42, 518-523. Hill, A. C. (1993). Mycoplasma indiense sp.nov., isolated from the throats of nonhuman primates. Int. J. Syst. Bacteriol. 43, 36-40. Hill, A. C , Polak-Vogelzang, A. A., and Angulo, A. F. (1992). Acholeplasma multilocale sp.nov., isolated from a horse and rabbit. Int. J. Syst. Bacteriol. 42, 513-517. International Committee on Systematic Bacteriology, Subcommittee on the Taxonomy of Mollicutes (1995). Revised minimum standards for description of new species of the class Mollicutes (Division Tenericutes). Int. J. Syst. Bacteriol. 45, 605-612. International Research Programme on Comparative Mycoplasmology (1990). "Report of Consultations." Int. Org. Mycoplasmol., Istanbul, Turkey. Jasper, D. E. (1994). Mycoplasmas and bovine mastitis. In " Mycoplasmosis in Animals: Laboratory Diagnosis" (H. W. Whitford, R. F. Rosenbusch, and L. H. Lauerman, eds.), pp. 62-67. Iowa State Univ. Press, Ames. Jordan, F. T. W. (1983). Recovery and identification of avian mycoplasmas. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 69-79. Academic Press, New York. Kahane, I., and Adoni, A., eds. (1993). "Rapid Diagnosis of Mycoplasmas." Plenum, New York.
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Kleven, S. H. (1994). Avian mycoplasmas. In "Mycoplasmosis in Animals: Laboratory Diagnosis" (H. W. Whitford, R. F. Rosenbusch, and L. H. Lauerman, eds.), pp. 31-38. Iowa State Univ. Press, Ames. Lauerman, L. H. (1994). Mycoplasmas of the bovine respiratory tract. In "Mycoplasmosis in Animals: Laboratory Diagnosis" (H. W. Whitford, R. F. Rosenbusch, and L. H. Lauerman, eds.), pp. 50-56. Iowa State Univ. Press, Ames. Maniloff, J., McElhaney, R. N., Finch, L. R., and Baseman, J. B., eds. (1992). "Mycoplasmas: Molecular Biology and Pathogenesis." Am. Soc. Microbiol., Washington, DC. McCabe, S. J., Murray, J. F., Ruhnke, H. L., and RachUs, A. (1987). Mycoplasma infection of the hand acquired from a cat. J. Hand Surg. 12A, 1085-1088. McCormack, W. M., Lee, Y. H., and Zinner, S. H. (1973). Sexual experience and urethral colonization with genital mycoplasmas: A study in normal men. Ann. Intern. Med. 78, 696-698. McCoy, R. E., Caudwell, A., Chang, C. J., Chen, T. A., Chiykowski, L. N., Cousin, M. T., Dale, J. L., de Leeuw, G. T. N., Golino, D. A., Hackett, K. J., Kirkpatrick, B. C , Marwitz, R., Petzold, H., Sinha, R. C , Sugiura, M., Whitcomb, R. F., Yang, I. L., Zhu, B. M., and Seemiiller, E. (1989). Plant diseases associated with mycoplasma-like organisms. In "The Mycoplasmas" (R. F. Whitcomb and J. G. TuUy, eds.). Vol. 5, pp. 545-640. Academic Press, San Diego, CA. Meyer, R. D., and Clough, W. (1993). Extragenital Mycoplasma hominis infections in adults: Emphasis on immunosuppression. Clin. Infect. Dis. 17(Suppl. 1), S243-S249. Ogata, S. (1983). Recovery and identification of canine and feline mycoplasmas. In "Methods in Mycoplasmology" (J. G. TuUy and S. Razin, eds.). Vol. 2, pp. 105-113. Academic Press, New York. Razin, S. (1991). The genera Mycoplasma, Ureaplasma, Acholeplasma, Anaeroplasma, and Asteroleplasma. In "The Prokaryotes" (A. Balows, H. G. Triiper, M. Dworkin, W. Harder, and K.-H. Schleifer, eds.), 2nd ed.. Vol. 2, 1937-1959. Springer-Verlag, New York. Razin, S. (1992). Mycoplasma taxonomy and ecology. In "Mycoplasmas: Molecular Biology and Pathogenesis" (J. Maniloff, R. N. McElhaney, L. R. Finch, and J. B. Baseman, eds.), pp. 3 22. Am. Soc. Microbiol., Washington, DC. Razin, S., and Barile, M. F., eds. (1985). "The Mycoplasmas," Vol. 4. Academic Press, New York. Razin, S., and Jacobs, E. (1992). Mycoplasma adhesion. J. Gen. Microbiol. 138, 407-422. Razin, S., and Tully, J. G., eds. (1983). "Methods in Mycoplasmology," Vol. 1. Academic Press, New York. Robinson, I. M., and Freundt, E. A. (1987). Proposal for an amended classification of anaerobic moUicutes. Int. J. Syst. Bacteriol. 37, 78-81. Rose, D. L., Tully, J. G., and Langford, E. V. (1978). Mycoplasma citelli, a new species from ground squirrels. Int. J. Syst. Bacteriol. 28, 567-572. Rosendal, S. (1994). Ovine and caprine mycoplasmas. In "Mycoplasmosis in Animals: Laboratory Diagnosis" (H. W. Whitford, R. F. Rosenbusch, and L. H. Lauerman, eds.), pp. 84-96. Iowa State Univ. Press, Ames. Ross, R. F., and Whittlestone, P. (1983). Recovery of, identification of, and serological response to porcine mycoplasmas. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 115-127. Academic Press, New York. Ruhnke, H. L. (1994). Mycoplasmas associated with bovine genital tract infections. In "Mycoplasmosis in Animals: Laboratory Diagnosis" (H. W. Whitford, R. F. Rosenbusch, and L. H. Lauerman, eds.), pp. 56-62. Iowa State Univ. Press, Ames. Ruhnke, H. L., and Madoff, S. (1992). Mycoplasma phocidae sp.nov., isolated from harbor seals (Phoca vitulina L). Int. J. Syst. Bacteriol. 42, 211-214. Salih, M. M., Friis, N. F., Arseculeratne, S. N., Freundt, E. A., and Christiansen, C. (1983). Mycoplasma mustelae, a new species from mink. Int. J. Syst. Bacteriol. 33, 476-479.
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Smith, P. F., and Morton, H. E. (1951). Isolation of pleuropneumonia-like organisms from the throats of humans. Science 113, 623-624. Somerson, N. L., and Cole, B.C. (1979). The mycoplasma flora of human and nonhuman primates. In "The Mycoplasmas" (J. G. Tully and R. F. Whitcomb, eds.). Vol. 2, pp. 191-216. Academic Press, New York. Taylor-Robinson, D., and Dinter, Z. (1968). Unexpected serotypes of mycoplasmas isolated from pigs. J. Gen. Microbiol. 53, 221-229. ter Laak, E. A., Tully, J. G., Noordergraaf, H. H., Rose, D. L., Carle, P., Bove, J. M., and Smits, M. A. (1993). Recognition of Mycoplasma canis as part of the mycoplasmal flora of the bovine respiratory tract. Vet. Microbiol. 34, 175-189. Tully, J. G. (1985). Newly discovered mollicutes. In "The Mycoplasmas" (S. Razin and M. F. Barile, eds.), Vol. 4, pp. 1-26. Academic Press, New York. Tully, J. G. (1989). Class Mollicutes: New perspectives from plant and arthropod studies. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.), Vol. 5, pp. 1-31. Academic Press, New York. Tully, J. G. (1993). Current status of the moUicute flora of humans. Clin. Infect. Dis. 17(Suppl.l), S2-S9. Tully, J. G., and Razin, S., eds. (1983). "Methods in Mycoplasmology," Vol. 2. Academic Press, New York. Tully, J. G., and Whitcomb, R. F., eds. (1979). "The Mycoplasmas," Vol. 2. Academic Press, New York. Tully, J. G., and Whitcomb, R. F. (1991). The genus Spiroplasma. In "The Prokaryotes" (A. Balows, H. G. Truper, M. Dworkin, W. Harder, and K.-H. Schleifer, eds.), 2nd ed., Vol. 2, pp. 1960-1980. Springer-Verlag, New York. Tully, J. G., Taylor-Robinson, D., Rose, D. L., Furr, P. M., and Hawkins, D. A. (1983). Evaluation of culture media for the recovery oi Mycoplasma hominis from the human urogenital tract. Sex. Transmit. Dis 10(Suppl.), 256-260. Tully, J. G., Rose, D. L., Clark, E., Carle, P., Bove, J. M., Henegar, R. B., Whitcomb, R. F., Colflesh, D. E., and Williamson, D. L. (1987). Revised group classification of the genus Spiroplasma (Class Mollicutes), with proposed new groups XII to XXIII. Int. J. Syst. Bacteriol. 37, 357-364. Tully, J. G., Whitcomb, R. F., Rose, D. L., Hackett, K. J., Clark, E., Henegar, R. B., Carle, P., and Bove, J. M. (1990). Current insight into the host diversity of acholeplasmas. Zentralbl. BakterioL, Suppl. 20, 461-467. Tully, J. G., Bove, J. M., Laigret, F., and Whitcomb, R. F. (1993). Revised taxonomy of the class Mollicutes: Proposed elevation of a monophyletic cluster of arthropod-associated mollicutes to ordinal rank (Entomoplasmatales, ord. nov.), with provision for familial rank to separate species with nonhelical morphology (Entomoplasmataceae, fam. nov.) from helical species (Spiroplasmataceae), and emended descriptions of the order Mycoplasmatales, family Mycoplasmataceae. Int. J. Syst. Bacteriol. 43, 378-385. Tully, J. G., Whitcomb, R. F., Rose, D. L., Bove, J. M., Carle, P., Somerson, N. L., Williamson, D. L., and Eden-Green, S. (1994a). Acholeplasma brassicae sp. nov., and Acholeplasma palmae sp. nov., two non-sterol-requiring mollicutes from plant surfaces. Int. J. Syst. Bacteriol. 44, 680-684. Tully, J. G., Whitcomb, R. F., Hackett, K. J., Rose, D. L., Henegar, R. B., Bove, J. M., Carle, P., Williamson, D. L., and Clark, T. B. (1994b). Taxonomic descriptions of eight new non-sterolrequiring mollicutes assigned to the genus Mesoplasma. Int. J. Syst. Bacteriol. 44, 685-693. Waites, K. B., Tully, J. G., Rose, D. L., Marriott, P. A., Davis, R. O., and Cassell, G. H. (1987). Isolation of Acholeplasma oculi from human amniotic fluid in early pregnancy. Curr. Microbiol. 15, 325-327.
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Whitcomb, R. F. and TuUy, J. G., eds. (1979). "The Mycoplasmas," Vol. 3. Academic Press, New York. Whitcomb, R. F. , and TuUy, J. G., eds. (1989). "The Mycoplasmas," Vol. 5. Academic Press, San Diego, CA. Whitford, H. W., and Lingsweiler, S. W. (1994). Canine, feline, and equine mycoplasmas. In "Mycoplasmosis in Animals: Laboratory Diagnosis" (H. W. Whitford, R. F. Rosenbusch, and L. H. Lauerman, eds.), pp. 134-140. Iowa State Univ. Press, Ames. Whitford, H. W., Rosenbusch, R. F., and Lauerman, L. H., eds. (1994). "Mycoplasmosis in Animals: Laboratory Diagnosis." Iowa State Univ. Press, Ames. Williamson, D. L., and Poulson, D. F. (1979). Sex-ratio organisms (spiroplasmas) of Drosophila. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.). Vol. 3, pp. 175-208. Academic Press, New York. Williamson, D. L., Tully, J. G., and Whitcomb, R. F. (1989). The genus Spiroplasma. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.). Vol. 5, pp. 71-111. Academic Press, San Diego, CA. Williamson, D. L., Tully, J. G., Rose, D. L., Hackett, K. J., Henegar, R., Carle, P., Bove, J. M., Colflesh, D. E., and Whitcomb, R. F. (1990). Mycoplasma somnilux, sp. nov.. Mycoplasma luminosum, sp. nov.. Mycoplasma lucivorax, sp. nov., sterol-requiring mollicutes from firefly beetles (Coleoptera; Lampyridae). Int. J. Syst. Bacteriol. 40, 160-164. Yechouron, A., Lefebvre, J., Robson, H. G., Rose, D. L., and Tully, J. G. (1992). Fatal septicemia with Mycoplasma arginini: A new human zoonosis. Clin. Infect. Dis. 15, 434-438.
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SECTION
A
Diagnostic Genetic Probes
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A1 INTRODUCTORY REMARKS Shmuel Razin
The new approach to rapid diagnosis of viral, bacterial, and parasitic infections, based on the development of gene or DNA probes, was introduced in the early 1980s. The principle is simple: genes or genomic sequences specific for a particular group of infectious agents, a single species, or even a particular strain are selected, cloned, or synthesized and used as probes hybridizing with DNA or RNA of the infected tissue or the clinical specimen. The more recent development of the polymerase chain reaction (PCR) enables the amplification of target DNA in the specimen, increasing the sensitivity of the tests by several orders of magnitude. Positive hybridization data, and in the case of PCR, demonstration in gels of the amplified target sequence, may suffice to indicate the presence of the infectious agent. The ultimate aim is to provide the laboratory with a diagnostic kit that will enable the detection and identification of the agent within a short time and with no need for special equipment. Although current probe tests are still more expensive than the traditional culture and biochemical tests, the rapid turnaround times of the new tests can translate into overall savings to the hospital and to the patient. The great interest in application of rapid diagnostic procedures to mycoplasma infections finds its expression in the proceedings of a workshop (Kahane and Adoni, 1993) and in a review devoted to this subject (Razin, 1994). The first DNA probes applied to diagnosis of mycoplasma infections consisted of cloned ribosomal RNA genes, such as the recombinant plasmid pMC5 (Amikam et al., 1982). The highly conserved rRNA genes were effective in detecting and identifying mycoplasmas infecting cell cultures by Southern blot hybridization with labeled pMC5 as a probe (Razin et al., 1984). In this way, 1 ng of mycoplasmal DNA, equivalent to about 10^ organisms, could be detected in contaminated cell cultures. Being highly conserved, the cloned rRNA gene 25 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Shmuel Razin
probes hybridize with the homologous genes of any mycoplasma and, in essence, with rRNA genes of any prokaryote. To provide more specific probes, synthetic oligonucleotides, 15 to 40 nucleotides in length, complementary to variable, species-specific regions of mycoplasmal 16S rRNA genes can be applied. When labeled, they can be used as probes in dot-blot hybridization with rRNA of the clinical specimens suspected of carrying the mycoplasma for which the probe was developed (Chapter A2, this volume). Since these probes hybridize to the mycoplasmal rRNA found in much more abundance than the DNA genomic targets, sensitivity of detection may reach 10^ to 10^ colony-forming units (CPUs), sufficient in most cases to detect mycoplasmas infecting cell cultures (Mattson and Johansson, 1993) but frequently insufficient to detect mycoplasmas in clinical samples, when small numbers of mycoplasmas are often present. Another class of DNA probes is that consisting of chromosomal segments specific for a certain mollicute species. These segments are derived from a genomic library of the specific mycoplasma, as described in Chapter A3 of this volume. The selected segments should not hybridize with DNAs of other mycoplasmas, particularly with those which may be present in the clinical specimen. Species-specific DNA probes, derived from genomic libraries, have been developed for a great variety of pathogenic mycoplasmas. Dot-blot hybridization with these probes, labeled either radioactively or by a variety of nonradioactive means (biotinylation, sulfonation, or by digoxigenin), exhibit a sensitivity level of 10^ to 10^ CPUs, a level which may not be sufficiently high for use in a clinical laboratory (Razin, 1994). The introduction of PCR in the late 1980s pushed aside many of the previously developed DNA probes and commercial kits. PCR tests are several orders of magnitude more sensitive than those based on direct hybridization with a DNA probe. Moreover, PCR is fast, copying a single DNA sequence over a billion times within 3 hours. Nucleic acid amplification techniques are not limited by the ability of an organism to grow in culture, a feature of paramount importance considering the fastidious nature of the mycoplasmas and the fact that not all mollicutes can be cultured in vitro, notably the phytoplasmas (see Chapter E6 in Vol. I, and D l l in this volume). PCR methodology is still at a phase of rapid development and new procedures and modifications are reported almost daily. Detailed descriptions of methodology and evaluation of PCR-based tests in diagnostics can be found in Rolfs et al. (1992) and in Persing et al. (1993). Chapters A4 to A8 (this volume) deal with the various aspects of the application of PCR methodology to diagnosis of mycoplasma infections. The first major issue in the development of a PCR-based test concerns the selection of the appropriate target sequences for amplification. The sequence to be amplified can be chosen from a published mycoplasmal gene sequence or from a randomly cloned DNA fragment demonstrated to be specific for the mycoplasma to be detected (Chapter A3, this volume). Complete or
Al Introductory Remarks
27
almost complete sequences of the 16S rRNA genes are now available for about 50 mollicute species and can be retrieved from several data banks (Chapter A2, this volume). This enables the selection of a variety of target sequences, starting with sequences in the highly conserved regions of the genes, producing primers of wide specificity ("universal primers") which will react with the DNA of any mycoplasma or even with the DNA of other prokaryotes. This may be satisfactory for detection of mycoplasma infection in cell cultures, where the goal is just to screen the cultures for contamination (Chapter F4, this volume). The mycoplasmal 16S rRNA genes carry, in addition to the conserved regions, more specific variable regions, as well as specific 16S-23S intergenic spacer regions. Primers can be selected from these regions with various degrees of specificity, ranging from clusters of species, single species, down to the subspecies level (Chapter A4, this volume). Specimen preparation for PCR testing is an important parameter that should be optimized. Since mycoplasmas have no cell walls, boiling of the sample, following its concentration by centrifugation, is often sufficient to make the organisms' DNA accessible. However, some clinical samples may contain undefined inhibitors of the PCR reaction, reducing the efficiency of amplification, so that DNA extraction has to be employed. Appropriate positive and negative controls should be included in each PCR test to rule out the presence of inhibitory substances (Chapters A5, A6, and F4, this volume). The methodology of target sequence amplification and optimization of the PCR test conditions as well as the identification of the PCR products are detailed in Chapter A6 (this volume). Although conventional PCR tests should, under optimal conditions, detect the DNA of a single mycoplasma, this rarely happens in testing clinical material. Two-step (nested) PCR has been devised to increase sensitivity to a level enabling detection of a single mycoplasma in a clinical sample. Another advantage of nested PCR is that the second-round PCR serves to confirm the specificity of the first-round PCR, as described and discussed in Chapter A7 (this volume). Another promising approach, described in Chapter A8 (this volume), is that termed random amplified polymorphic DNA or arbitrary primer PCR. It involves PCR amplification with a single arbitrary primer at low stringency, resulting in strain-specific arrays of DNA fragments that can reproducibly distinguish even closely related strains of a species.
References Amikam, D., Razin, S., and Glaser, G. (1982). Ribosomal RNA genes in mycoplasmas. Nucleic Acids Res. 10, 4215-4222. Kahane, I., and Adoni, A., eds. (1993). "Rapid Diagnosis of Mycoplasmas." Plenum, New York and London.
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Mattson, J. G., and Johansson, K.-E. (1993). Oligonucleotide probes complementary to 16S rRNA for rapid detection of mycoplasma contamination in cell cultures. FEMS Microbiol. Lett. 107, 139-144. Persing, D. H., Smith, T. F., Tenover, F. C , and White, T. J. (1993). "Diagnostic Molecular Microbiology: Principles and Applications." Am. Soc. Microbiol, Washington, DC. Razin, S. (1994). DNA probes and PCR in the diagnosis of mycoplasma infections. Mol. Cell. Probes %,A91-5n. Razin, S., Gross, M., Wormser, M., Pollack, Y., and Glaser, G. (1984). Detection of mycoplasmas infecting cell cultures by DNA hybridization. In Vitro 20, 404-408. Rolfs, A., Schuller, I., Finckh, U., and Weber-Rolfs, I. (1992). "PCR: Clinical Diagnostics and Research." Springer-Verlag, Berlin.
A2 OLIGONUCLEOTIDE PROBES COMPLEMENTARY TO 16S rRNA Karl-Erik Johansson
Introduction The ribosome is an essential organelle in all living cells as it constitutes the protein-synthesizing machinery. It is built up of proteins and ribosomal RNA (rRNA), and there are three species of rRNA in bacteria; 5S, 16S, and 23S rRNA. These molecules are about 120, 1500, and 3000 nucleotides in length and they have sequence regions of different evolutionary variability. A model of the secondary structure of the Escherichia coli 16S rRNA is shown in Fig. 1 and the regions of different evolutionary variability as defined by Gray et al., (1984) are also indicated. The localization and size of these regions are essentially the same in all bacteria. The possibility of using group-specific DNA probes complementary to ribosomal RNA, the so-called rDNA probes, for detection of mollicutes was first brought up by Stanbridge (1976). It was later applied to detect mycoplasma contamination in cell cultures by filter hybridization experiments with restriction fragments of rRNA genes as probes (Gobel and Stanbridge, 1984; Razin etal., 1984; Johansson etal., 1990). Today, synthetic oligonucleotides of 15-40 nucleotides in length are more commonly used as probes since the most suitable target region can be chosen, which in turn will result in the desired specificity (Gobel, 1991; Johansson, 1993). At least five factors are important in the utilization of rRNA as a target molecule for DNA probes. First, rRNA molecules are present in several thousand ribosomes in a growing bacterium, and a detection system based on hybrid29 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
30
Karl-Erik Johansson
Fig. 1 . Secondary structure model of 16S rRNA from Escherichia coli based on the model published by Woese et a/. (1983) and adapted from Gray et al. (1984) by Johansson (1993). Reproduced with kind permission from Plenum Press. Variable (V) regions (---), semiconserved (S) regions (—), and universal (U) regions . The position of each 100*^^ nucleotide is indicated with an arrow. Only the two S regions for which probes have been designed (see Table I) are indicated (SI a and S5b).
ization to rRNA will, therefore, be very sensitive. Second, the rRNA nucleotide sequence composed of regions of different evolutionary variability provides predestinate specificity. Species-specific as well as group-specific probes can be designed. Third, sequence analysis of 16S rRNA has proved to be an indispensable tool for evolutionary and phylogenetic studies, and the interest in using sequence data of rRNA for phylogenetic studies of mycoplasmas (Weisburg et
A2 rDNA Oligonucleotide Probes
31
a/., 1989) is great. Sequence data from rRNA are, therefore, rapidly accumulating in the databases (Olsen et al., 1992); this sequence information is invaluable for the design of rDNA probes. Fourth, it is comparatively easy to determine the complete sequence of the 16S rRNA molecule (or its gene) because of the presence of eight evenly distributed conserved regions of 50 to 110 nucleotides in length (Gray et al, 1984). These so-called universal regions (U1-U8) can be used as targets for sequencing primers or as targets for polymerase chain reaction (PCR) primers. Biotinylated PCR products can be produced for automated solidphase sequencing by which sequence data of rRNA genes can be generated very efficiently (Pettersson etal, 1994). Fifth, an identification system (hybridization or in vitro amplification by PCR) based on rRNA or rRNA genes is very robust, and a system by which one species is detected can most probably also be used for detection of all strains or subspecies within that species, provided that the target region has been properly chosen. This could, however, also be a drawback when it is essential to be able to differentiate between subspecies or variant strains.
Materials Design and Construction of Oligonucleotides Access to a computer with software for sequence analysis including the databases (EMBL data bank or GenBank) for nucleotide sequences greatly facilitates the design of rDNA probes. Suitable program packages, including GenBank on CD-ROM, for computers (IBM-compatible PCs, Macintosh, or SUN) with different operating systems (MS-DOS, Macintosh System, or UNIX) are available from IntelliGenetics (Mountain View, CA). Another program package for sequence analysis with a VAX computer (VMS operating system) is available from the Genetics Computer Group at the University of Wisconsin (Madison, WI) (Devereux et al., 1984). Other useful programs are also available from different companies. The oligonucleotide can be synthesized on a DNA synthesizer and different machines are available on the market. However, if only a limited number of oligonucleotides are constructed each year, it is probably more cost effective to order the oligonucleotide from a company which performs oligonucleotide synthesis on commission. Several such companies exist and their addresses can be found in advertisements in journals such as Science and Nature. Labeling of Oligonucleotides Radiolabeling is probably the first choice when starting to work with DNA probes; this is treated in detail in the following discussion. Radiolabeling is also
32
Karl-Erik Johansson
the most sensitive method and radiolabeled DNA probes do not exhibit nonspecific binding to the sample material, as is sometimes the case with nonradioactively labeled probes, unless preparations of purified nucleic acids or PCR products are applied onto the filters. However, in the long run it is of course highly desirable to be able to use nonradioactive labeling and different systems are available on the market. These systems are supplied as complete kits with detailed instructions. One commonly used system is based on the labeling of the oligonucleotide at the 3' terminus with digoxigenin-dideoxy(dd)UTP using terminal deoxynucleotidyltransferase (TDT). Digoxigenin is a steroid hapten to which antibodies can easily be raised. Hybridization signals are then obtained with such antibodies conjugated to alkaline phosphatase (AP) by an AP-catalyzed color reaction. For higher sensitivity, a tail of digoxigenin-deoxy(d)UTP can be coupled to the 3' terminus. Another system is based on the detection of hybridization signals by enhanced chemiluminescence (ECL). In this ECL system, the oligonucleotide is labeled at the 3' terminus with a tail of six to eight fluorescein-11-dUTP molecules using terminal deoxynucleotidyltransferase. Fluorescein is used as a hapten in this system also. Antibodies to fluorescein, conjugated with horseradish peroxidase (HRP), are added after hybridization and the HRP-modified oligonucleotide, bound to its target, can then be used to oxidize luminol in the presence of hydrogen peroxide which will result in light emission. The light emission can be registered on a special film. A. RADIOLABELING
Access to a laboratory where ^^P can be handled safely Standard pipettes Microcentrifuge Microcentrifuge tubes T4 polynucleotide kinase [7-32p]ATP (Amersham, Little Chalfont, UK, or DuPont, NEN Research Products, Boston, MA) NAP-5 columns (Pharmacia LKB Biotechnology, Uppsala, Sweden) Thermostatted water bath Scintillation counter Kinase buffer (stock solution, 10x): 500 mM Tris-HCl (pH 7.4) 100 mM MgCl2 50 mM dithiothreitol (DTT) 10 mM spermidine (optional) Elution buffer (=TENS buffer): 100 mM Tris-HCl (pH 8.0) 100 mM NaCl
A2 rDNA Oligonucleotide Probes
33
1 mM EDTA 0.5% Sodium dodecyl sulfate (SDS) B. LABELING WITH DIGOXIGENIN-ddUTP AND DETECTION BY AN ENZYMECATALYZED COLOR REACTION
DIG oligonucleotide 3' end labeling kit (Boehringer Mannheim Biochemica, Mannheim, Germany) Positively charged nylon membrane from Boehringer Mannheim is recommended by the manufacturer C. LABELING WITH FLUORESCEIN-11-dUTP AND DETECTION BY ECL
ECL 3'-oligolabeling and detection system (Amersham) The positively charged nylon membrane Hybond-N+ (Amersham) is recommended by the manufacturer Hyperfilm ECL (Amersham) is recommended by the manufacturer
Filter Hybridization Standard pipettes Microcentrifuge Microcentrifuge tubes Dot-blot filtration manifold (from Bio-Rad Laboratories, Richmond, CA or Schleicher & Schuell GmbH, Dassel, Germany) Blotting medium (Hybond C Extra from Amersham or Zeta probe GT from BioRad) ELISA microtiter plates (for dilution of samples) Hybridization oven equipped with a rotisserie Glass vials to be used in the hybridization oven Thermostatted waterbath X-ray films (X-OMAT AR film from Eastman Kodak Co., Rochester, NY) X-ray film cassettes with intensifying screens (Cronex Lightning Plus from DuPont, NEN Research Products) UV cross-linker (Amersham or Stratagene, La Jolla, CA) (optional) Automatic X-ray film developer (optional) X-ray developer (Kodak D19) Fixative Phosphate-buffered saline, pH 7.4 (PBS) SDS (20% stock solution)
34
Karl-Erik Johansson
Frozen (—70°C) aliquots of reference cultures of appropriate mycoplasmas with known CFU (colony-forming units) [or CCU (color-changing units)]. Hybridization solution: Denhardt's solution (50 x) SSC (20 X) Herring (or salmon) sperm DNA (10 mg/ml) SDS (20%)
Hybond C Extra 2.00 ml 1.00 ml 0.15 ml
— 6.85 ml
Zeta Probe GT 2.00 1.00 0.15 1.00 5.85
ml ml ml ml ml
Prepare the herring (salmon) sperm DNA solution as described by Sambrook etal. (1989). Denhardt's solution (50x): 5 g Ficoll (Type 400, Pharmacia Biotech.) 5 g polyvinylpyrrolidone, average M^. 360,000 (Sigma Chemical Co., St. Louis, MO) 5 g bovine serum albumin (Fraction V from Sigma Chemical Co.) Dissolve the components in autoclaved water to a final volume of 500 ml, filter the solution, dispense into aliquots, and store at -20°C. SSC (20X): 3.0 MNaCl (175.3 g) 0.3 M sodium citrate (C6H5Na307-2H20) (82.2 g) H2O (800 ml) Adjust the pH to 7.0 and the volume to 1000 ml. Autoclave the solution.
Procedure Design and Construction of Oligonucleotides Several group- and species-specific rDNA oligonucleotide probes have been designed for the detection and identification of mycoplasmas. Some of the properties of these probes and their sequences are listed in Tables I and II, respectively. The important considerations for the design of such probes are discussed next. 1. It is of course essential to have sequence information of the 16S rRNA molecule of the species for which the detection system is to be developed. If the sequence is not available in the databases (GenBank or EMBL) or in the literature, the sequence of the rRNA molecule or its gene must first be determined with sequencing primers complementary to universal regions (see Chapter E3,
A2 rDNA Oligonucleotide Probes
35
Vol. I). It is sometimes sufficient to make a partial sequence determination. However, it is always recommended to determine the complete sequence since it is easier to design an optimal system when the complete sequence is known. 2. The sequence should then be compared by computer alignment with the corresponding sequences of primarily closely related species which are likely to be found in the same ecological niche. 3. The next step is to select a suitable target region with as many sequence differences as possible compared to the corresponding region from the species for which it is essential to avoid cross-hybridization. These sequence differences should be evenly distributed, but are less efficient when close to the termini of the probe. It has been shown in some cases and under optimal conditions (high stringency) that one single mismatched base pair is sufficient to discriminate between a homologous and a heterologous target when the probe is short (about 15 nucleotides). It is important to have a high G+C content (about 40%) which gives stronger hybridization signals. Some high quality (unstable) mismatches should also be included, like A-A, U-U, C-U, or C-A, which will decrease the risk of cross-hybridization between the probe and the heterologous targets (Mattsson etaL, 1991). When a tentative target region has been selected, it could also be valuable to make a global search in the database for the selected target region to check if there are other species to which the probe might cross-hybridize. The target region for a species-specific rDNA probe will most probably be a segment of one of the eight evolutionarily variable (V) regions (cf. Fig. 1 and Table I). 4. If no serious cross-hybridizations can be anticipated, the corresponding oligonucleotide can be synthesized or ordered. Note that the nucleotide sequence of the oligonucleotide should be complementary to and reversed of the target region. Different modifications of the oligonucleotide can be introduced during synthesis and these modifications can then be utilized for nonradioactive labeling or detection. 5. The next step is to test the specificity and sensitivity of the probe by filter hybridization experiments with reference cultures of mycoplasmas (see later). It is important to include relevant species for which sequence information is not available. Labeling of Oligonucleotides Detailed instructions for the nonradioactive labeling of oligonucleotides are provided with the kits used for that purpose and are, therefore, not described here. Oligonucleotides can be radiolabeled at the 5' terminus with T4 polynucleotide kinase (T4 PNK) and [7-^2p]A'rP as the label or at the 3' terminus with TDT and [a-^^pj^^jsjxP as the label. When TDT is used, it is also possible to add a tail of labeled nucleotides to the 3' terminus for higher sensitivity by using a deoxynucleotide instead of a dideoxynucleotide. Labeling with T4 PNK and TDT is described in many manuals in molecular biology (Sambrook et al., 1989)
TABLE I
Designation
Regionb
Length
TmC("c)
Specificity
Known cross-hybridizations
Refs.
--
MYC14 MYC25
U2 Sla
14 25
34 69
Mollicutes Mollicutes
MP20 MP30
V6 V3
20 30
58 72
Mycoplasma pneumoniae M. pneumoniae
Mbg28
V2
28
70
M. bovigenitalium
Mbo29 Mag30
V6 V8
29 30
69 70
M. bovis M. agalactiae
Gobel et al. (1987a) Gobel et al. (1987a)
M. genitalium
G8bel er al. (1987b) Gobel et al. (1987b)
C. FemBndez, J. G. Mattsson, U. B. Gobel, and K. E. Johansson (personal communication) M. agalactiae M . bovis, M. bovigenitalium, M . californicum
Mattsson et al. (1991) Mattsson et a / . (1991)
Mhp8124 Mhp3132 Mhp6130 Mhy2127 MHPl MHP2 MHP3
34 69 58 72 70
hyopneumoniae hyopneumoniae hyopneumoniae hyorhinis
M . hyopneumoniae M . hyopneumoniae M . hyopneumoniae M . gallisepticum M . synoviae
Mga V8131 Msy V8131 Mcc l M C C ~ ~ M c c ~ ~
M. M. M. M.
69 70
M . jlocculare M . jlocculare
M . jlocculare M . jlocculare
M . imitans
Hominis groupe Acholeplasma laidlawii M. pneumoniae clusterr
.The sequences of the probes (except MYC14 and MYC25) are given in Table 11. hTarget region of 16s rRNA according to the nomenclature of Gray et al. (1984). cMelting temperature of the corresponding DNA-DNA homoduplex in 2 X SSC. dA mixture of Mccl, Mcc2, and Mcc3 can be used to detect most mycoplasmas found as cell culture contaminants.
Johansson Johansson Johansson Johansson
et al. et al. et al. et al.
(1992) (1992) (1992) ( 1992)
Futo et al. (1992) Futo et al. (1992) Futo et al. (1992) Fernandez et al. (1993) Fernandez et al. (1993) Mattsson and Johansson (1993) Mattsson and Johansson (1993) Mattsson and Johansson (1993)
38
Karl-Erik Johansson
TABLE 11 SEQUENCES OF rDNA OLIGONUCLEOTIDE PROBES FOR DETECTION AND IDENTIFICATION OF MYCOPLASMAS^
Designation
Sequence
MP20 MP30
5'-CTCTAGCCAT TACCTGCTAA-3' 5'-CCACCTGTCA CTCGGTTAAC CTCCATTATG-3'
Mbg28
5'-TTCCGTAAAT GCCATGCGGC ATCTACGA-3'
Mbo29 Mag30
5'-CGTCAAGGTA GCATCATTTC CTATGCTAC-3' 5'-TGCGTCGATG AGTTCCCCAT CAACTAATGA-3'
Mhp8/24 Mhp3/32 Mhp6/30 Mhy2/27
5'-TGTGTTAGTG ACTTTTGCCA CCAA-3' 5'-TGATCTCGTT AGCCTCGGCT ATATCTCTAT AG-3' 5'-CCGTCAAGAC TAGAGCATTT CCTATCTAGT-3' 5'-CTATTACTCA TCATGCGATA AATAACT-3'
MHPl MHP2 MHP3
5'-CTTGGTGAAG CTTGAAGGCT CCTTTGAATA-3' 5'-CGGACTAAAG TTGAGCTTTA GCATTTAACT-3' 5'-TAGCCTCGGC TATATCTCTA TAGTTTTGCG-3'
Mga V8/31 Msy V8/31
S'-ACTGCAGCAC CGAAGTATTC GCTCCGACAC T-3' 5'-GCTGCGTCGA TGGTTTCCAT CAACTAGTCA T-3'
Mccl Mcc2 Mcc3
5'-GCCCCACTCG TAAGAGGCAT G-3' 5'-TAAGCACTGC ACTTACACCA C-3' 5'-GCACGTTTGC AGCCCTAGAC A-3'
^References to the literature and the properties of the probes are listed in Table I.
and kits are also available for radiolabeling with these enzymes. However, the former method will also be described here since it is one of the most commonly used procedures for labeling oligonucleotides. It is important to follow the local regulations concerning handling of radioactive material. Keep the radioactive stock solutions in lead containers, work behind a plexiscreen (1.5 cm in thickness), and use protective glasses. Use autoclaved tips and tubes throughout. 1. Add the following components to a microcentrifuge tube: H2O Kinase buffer (lOx) DNA (about 200 ng) T4 PNK (10 U/M-1) [T-32p]ATP (1.85 MBq, >185 MBq/mmol) Final volume
. . . . ^,1 2.0 \j.\ . . . . M-l 0.7 |xl 5.0 \i\ 20.0 jxl
A2 rDNA Oligonucleotide Probes
39
The amount of water is dependent on the concentration of the probe (DNA). A suitable buffer (for instance, One-Phor-AU buffer PLUS from Pharmacia Biotech.) is often supplied with the enzyme. Spin for a few seconds to mix and incubate at 37°C for 45 minutes. Seal the tube with a clip and incubate at 65°C for 5 minutes to inactivate the enzyme. Spin the tube again and add 80 jxl of elution buffer. 2. Remove the excess of radioactive ATP by gel filtration on a NAP-5 column equilibrated with elution buffer. Work behind the plexiscreen. Apply the sample onto the gel bed and elute it into the column with 400 |xl of elution buffer. Collect the first 500 jxl which correspond to the void volume of the column in one tube. Add 3 x 330 |xl of elution buffer and collect the three corresponding fractions in three tubes. Store the fractions in a shielded container. 3. Determine the radioactivity in 5 |xl (diluted into, for instance, 100 |xl of elution buffer) from the three last fractions with a scintillation counter by utilizing the Cerenkov effect in the tritium channel or with a fully opened window. The efficiency is approximately 55%. Select the fraction with the highest radioactivity for maximal sensitivity. This fraction should contain 2-5 x 10^ cpm/5 |JL1 of probe solution in a successful labeling experiment. Store the labeled oligonucleotide at — 20°C and use it within 1 week for optimal sensitivity. The oligonucleotide is sensitive to radiolysis.
Sample Preparation and Application onto Membrane Filters Both Hybond C Extra, which is a nylon-reinforced nitrocellulose membrane, and Zeta Probe GT, which is a pure nylon membrane, are mechanically very stable. Membrane filters should, however, always be handled with care. Use gloves or tweezers (with flat ends) to avoid getting grease on the membrane. Use a soft pencil and a clean ruler to mark where to cut the membrane. 1. Cut the membrane filter to a suitable size and pre wet it in PBS before use. 2. Centrifuge 1 ml of mycoplasma culture (about 10^ CFU/ml) at about 12,000 g for 5-10 minutes and suspend the pellet in 1 ml of PBS. Keep the suspension on ice. 3. Load the dot-blot filtration manifold with a membrane filter according to the instructions of the manufacturer. Apply about 250 JJLI of the undiluted suspension and in a suitable dilution series onto the membrane filter. The samples can be conveniently diluted in an ELISA microtiter plate with an 8- or 12-channel pipette. Be careful to avoid cross-contamination. Wash the dot-blot manifold carefully after use to avoid contamination problems in the next experiment. 4. Mycoplasmas are easily lysed and the rRNA can, therefore, readily be made accessible for hybridization. When radiolabeled probes are used in the hybridization experiments, it is not necessary to isolate RNA from the sample. It
40
Karl-Erik Johansson
can be applied directly onto the filter and the mycoplasmas are then lysed on the filter (see later). 5. Prepare some extra membranes. One of them should be used for hybridization with a general probe (complementary to a universal region). This is an important control experiment to check that the same amounts of mycoplasmas were applied onto the filter, or rather that the same amounts of rRNA are accessible for hybridization with the probe. The amount of sample applied onto the filter should be adjusted to give the same signal strength with a general probe before trying the specific probe. This is very important for correct interpretation of the results because the number of ribosomes is dependent on the growth of the cell and can vary considerably. The following oligonucleotide can be labeled and used as a general probe for mycoplasmas (and many other gram-positive bacteria): 5'-CCGTCAATTC MTTTYAGTTT-3' (Lane et aL, 1985). This oligonucleotide is complementary to the U5 region and should be synthesized with two ambiguities for maximal generality. The melting temperature (see later) for the "U5 probe" is 37°-4rC. 6. When samples containing more or less intact mycoplasmas have been applied onto the filter for direct filter hybridization, lysis of the cells, denaturation, and fixation of nucleic acid can be achieved by boiling the membrane for 5 minutes in about 500 ml of a solution containing 0.01 x SSC and 0.01% SDS (Johansson et al., 1990). After drying, the membrane can be used directly in the hybridization experiment with radiolabeled probes (Fig. 2A). Nonradioactively labeled probes are better suited for hybridization to samples consisting of pure nucleic acid preparations or PCR products (Fig. 2B). The PCR products have to be transformed to single-stranded DNA by heating the sample in a total volume of 0.5 ml of 0.4 M NaOH and 10 mM EDTA at 100°C for 10 minutes. The samples can then be applied onto the membrane filter. The membrane should then be taken out from the apparatus and rinsed briefly in 2x SSC to neutralize the samples before air drying. Fixation of DNA preparations and PCR products onto nylon membranes can be achieved by cross-linking under UV light.
Hybridization Hybridization is the reaction by which a nucleic acid probe binds to a complementary target molecule by base pairing. The target molecule can be DNA or RNA. The prehybridization and hybridization steps can be performed statically in heat-sealable plastic bags, but it is more convenient and faster to use a hybridization oven for incubation under agitation in the rotisserie. 1. Estimate the melting temperature (T^) for the oligonucleotide and its target in a hypothetical DNA-DNA homoduplex according to the following formula if the oligonucleotide is short (<18 nucleotides):
41
A2 rDNA Oligonucleotide Probes
T^ = 4(G + C) + 2(A + T). The following more sophisticated formula can be used for oligonucleotide probes up to 100 nucleotides in length: T^ = 16.6 log[Na+] + 0.41Pg, + 81.5 - P^^ - 675/L - 0.65Pf,, where P^^ is the G + C content in % (should be 30-70%); P^^ is the number of mismatched base pairs in %; P^^ is [formamide] in %; and L is the length of the oligonucleotide in number of nucleotides. This formula can also be used to calculate T^ when a certain number of mismatches can be tolerated. When long oligonucleotides (with a high G + C content) are used, it might be necessary to decrease T^ by including formamide during hybridization. Perform the hybridization at about 5°C under T^. Try,
1
2
3
4
B 3
4
B
C D i
. l
l
# ^<
E F Fig. 2. Filter hybridization of ampiicons from porcine mycoplasmas obtained by in vitro amplification by PCR with primers complementary to the V2 and the V8 regions of the 16S rRNA gene of Mycoplasma hyopneumoniae (Mattsson et a/., 1995). The filter was hybridized with the M h p 6 / 3 0 probe (see Tables I and II) labeled with (A) 32p and (B) digoxigenin-ddUTP. Hybridization signals were obtained by (A) autoradiography to an ordinary X-ray film and (B) an enzyme-catalyzed color reaction with alkaline phosphatase-conjugated antibodies to digoxigenin. The following samples were applied in a 10-fold dilution series: A - B 1 - 2 , M. hyorhinis; C - D 1 - 2 , M. hyosynoviae; E-F 1-2, M. hyopharyngis; A - B 3 - 4 , M. sualvi; C - D 3 4, M. hyopneumoniae; E-F 3 - 4 , M. arginini. Columns 1 and 3 represent 20 |JLI of undiluted PCR product, and the samples applied in columns 2 and 4 contain 2 |JLI. Each sample was analyzed in duplicate (PCR from 5 jxl undiluted and 5 ^JLI of 10-fold-diluted samples applied in the upper and lower row, respectively).
42
Karl-Erik Johansson
however, different hybridization temperatures (T{) for optimal results because it is difficult to calculate T^ for a DNA-RNA heteroduplex which is significantly more stable than a DNA-DNA homoduplex. 2. Prehybridize for at least 30 minutes at T^ under agitation in a glass vial suitable for the hybridization oven. 3. Add about 2 x 1 0 ^ cpm of the probe per milliliter of hybridization solution and continue the incubation for 2-3 hours. Use 5-10 ml of hybridization solution for incubation in the glass vials. Smaller volumes (and a corresponding higher activity) can be used if the filter is incubated statically in a heat-sealable plastic bag since the size of the bag can be adapted to the filter. Note that SDS has to be added to the hybridization solution to avoid high background if Zeta Probe GT is used as a blotting medium. 4. Rinse the filter under agitation in 50-200 ml of SSC (2x) for 5 minutes at Ti and for 3 x 20 minutes at Tj-lO^C. The smaller volume has to be used in the glass vials for the hybridization oven and the larger volume can be used if plastic vessels are utilized. It is very important to prewarm the SSC solution to the correct temperature before starting the rinsing process, particularly for the first 5-minute wash at Tj. 5. Dry the filter, affix it with tape to a piece of stiff filter paper, and wrap it in thin plastic foil. Cut the upper left comer for easy orientation in the dark room and make a corresponding cut in the X-ray film. Expose the film overnight at —70°C in a cassette for X-ray films equipped with an intensifying screen for P-radiation. The filter should not be completely dried if it is to be reprobed. In this case, put the wet filter in a heat-sealable plastic bag for exposure. Such a filter can be erased by boiling as described earlier under Sample Preparation and Application onto Membrane Filters, step 6, and reprobed. 6. Develop the film for 3 minutes in an X-ray developer, give the film a quick rinse in water, put it in the fixative for 3 minutes, and wash it again for 30 minutes. Dry the film. A machine for automatic development of X-ray films is very convenient, but it has to be used daily for optimal performance and is, therefore, only practical in a large laboratory. 7. Evaluate the result. If unwanted cross-hybridizations are obtained, it is necessary to increase the stringency of the system by increasing the temperature of hybridization and/or washing. The hybridization and/or the washing can also be performed at lower salt concentrations which will also increase the stringency. If cross-hybridizations persist, it might be necessary to choose another target region for the probe. A useful, but very expensive, alternative to autoradiography with X-ray films is to utilize a system for reading reusable image screens. Such a screen is covered by a layer of BaFBr:Eu2+ crystals which can be activated by ionizing radiation from, for instance, a membrane filter. The activated screen is then read by
A2 rDNA Oligonucleotide Probes
43
scanning with a laser beam in an image reading unit in which the emitted energy can be registered. The screen can then be completely erased by exposure to strong light and used again. Several systems are available on the market, for instance, the Phosphorimager (Molecular Dynamics, Sunnyvale, CA) and Fujix BAS 2000 (Fuji Photo Film Co., Tokyo, Japan). A system based on laser scanning of reusable image screens is very convenient since it is quick (exposure for 2-3 hours corresponds to an overnight exposure to an ordinary X-ray film for ^2p), it has a broad dynamic linear range, and it is quantitative. Furthermore, the system is completely computerized and printouts ready for publishing can be obtained immediately (see Fig. 3).
Discussion The expectations from use of rDNA probes in direct filter hybridization of clinical material were high at the beginning. However, despite the high sensitivity of a diagnostic system based on rDNA probes, it is still not sensitive enough for many clinical applications and it is difficult to analyze the kind of samples that can be collected from live animals. However, infected tissue can be analyzed, and Mycoplasma hyopneumoniae has been detected in lung tissue from experimentally infected pigs by direct filter hybridization with the Mhp6/30 probe (Johansson et al., 1992). It would be highly desirable, however, to be able to detect the organisms in, for instance, nose swabs from pigs, but the sensitivity of the probe is not sufficient for that purpose. One application where rDNA oligonucleotide probes have proved to be useful for direct filter hybridization of samples is for mycoplasma screening of cell cultures (Mattsson and Johansson, 1993). Mycoplasmas are present in large numbers in infected cell lines and the sensitivity of a probe-based detection system will, therefore, be sufficient in most cases. Probes can also be useful for species identification of mycoplasmas grown in culture where the amount of mycoplasmas present in the sample is, in general, not a limiting factor. Several species-specific rDNA oligonucleotide probes have been designed for mycoplasmas (Tables I and II). Another important use is for confirmation of PCR experiments (see Chapters A4-A8 in this volume) illustrated in Fig. 2. Radiolabeled probes can be used for this purpose (Van Kuppeveld et al, 1992), but it is also possible to use nonradioactively labeled probes since a very large number of target molecules can be applied onto the filter after in vitro amplification by PCR. Furthermore, since the material applied onto the filter is essentially pure DNA, the risk of interference with components in the clinical material is negligible. This is illustrated in Fig. 2B which shows a filter hybridization experiment with the rDNA probe Mhp6/30
44
Karl-Erik Johansson
-mr
TOG
»«8yi«|j
"TO:!
-TCI
"wri
-in
WE^wi:^
s: "wm m:m "Oif Ira
TM
A.
UP
Fig. 3. Autoradiography of the filter shown in Fig. 2A to a reusable image screen and laser beam scanning with a Fujix BAS 2000. Spots 1 - 4 correspond to C-D 3-4 (the PCR products of M. hyopneumoniae) in Fig. 2. Spot 5 was used to determine the background radiation. The radioactivities in spots 1-4 are listed in the tabulation.
A2 rDNA Oligonucleotide Probes
45
(Tables I and II) labeled with digoxigenin-ddUTP. PCR products from a segment of the 16S rRNA gene from different porcine mycoplasmas were used as samples. The same probe labeled with ^^p was used for comparison (Fig. 2A).
References Devereux, J., Haeberli, P., and Smithies, O. (1984). A comprehensive set of sequence analysis programs for the VAX. Nucleic Acids Res. 12, 387-395. Fernandez, C , Mattsson, J. G., Bolske, G., Levisohn, S., and Johansson, K.-E. (1993). Speciesspecific oligonucleotide probes complementary to 16S rRNA of Mycoplasma gallisepticum and Mycoplasma synoviae. Res. Vet. Sci. 55, 130-136. Futo, S., Seto, Y., Mitsuse, S., and Mori, Y. (1992). Detection of Mycoplasma hyopneumoniae by using rRNA-oligodeoxynucleotide hybridization. J. Clin. Microbiol. 30, 1509-1513. Gobel, U. B. (1991). Targeting ribosomal RNA sequences: A universal approach to the detection and identification of microorganisms. In "Rapid Methods and Automation in Microbiology and Immunology" (A. Vaheri, R. C. Tilton, and A. Balows, eds.), pp. 27-36. Springer-Verlag, Berlin. Gobel, U. B., and Stanbridge, E. J. (1984). Cloned mycoplasma ribosomal RNA genes for the detection of mycoplasma contamination in tissue cultures. Science 116^ 1211-1213. Gobel, U. B., Maas, R., Haun, G., Vinga-Martins, C , and Stanbridge, E. J. (1987a). Synthetic oligonucleotide probes complementary to rRNA for group- and species-specific detection of mycoplasmas. Isr. J. Med. Sci. 23, 742-746. Gobel, U. B., Geiser, A., and Stanbridge, E. J. (1987b). Oligonucleotide probes complementary to variable regions of ribosomal RNA discriminate between Mycoplasma species. J. Gen. Microbiol. 133, 1969-1974. Gray, M. W., Sankoff, D., and Cedergren, R. J. (1984). On the evolutionary descent of organisms and organelles: A global phylogeny based on a highly conserved structural core in small subunit ribosomal RNA. Nucleic Acids Res. 12, 5837-5852. Johansson, K.-E. (1993). Detection and identification of mycoplasmas with diagnostic DNA probes complementary to ribosomal RNA. In "Rapid Diagnosis of Mycoplasmas" (I. Kahane and A. Adoni, eds.), pp. 139-154. Plenum, New York. Johansson, K.-E., Johansson, I., and Gobel, U. B. (1990). Evaluation of different hybridization procedures for the detection of mycoplasma contamination in cell cultures. Mol. Cell. Probes 4, 33-42. Johansson, K.-E., Mattsson, J. G., Jacobsson, K., Fernandez, C , Bergstrom, K., Bolske, G., Wallgren, P., and Gobel, U. B. (1992). Specificity of oligonucleotide probes complementary to evolutionarily variable regions of 16S rRNA from Mycoplasma hyopneumoniae and Mycoplasma hyorhinis. Res. Vet. Sci. 52, 195-204. Lane, D. J., Pace, B., Olsen, G. J., Stahl, D., Sogin, M. L., and Pace, N. R. (1985). Rapid determination of 16S ribosomal RNA sequences for phylogenetic analysis. Proc. Natl. Acad. Sci. USA 82, 6955-6959. Mattsson, J. G., and Johansson, K.-E. (1993). Oligonucleotide probes complementary to 16S rRNA for rapid detection of mycoplasma contamination in cell cultures. FEMS Microbiol. Lett. 107, 139-144. Mattsson, J. G., Gersdorf, H., Gobel, U. B., and Johansson, K.-E. (1991). Detection of Mycoplasma bovis and Mycoplasma agalactiae by oligonucleotide probes complementary to 16S rRNA. Mol. Cell. Probes 5, 27-35.
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Karl-Erik Johansson
Mattsson, J. G., Bergstrom, K., Wallgren, P., and Johansson, K.-E. (1995). Detection of Mycoplasma hyopneumoniae in nose swabs from pigs by in vitro amplification of the 16S rRNA gene. J. Clin. Microbiol. 33, 893-897. Olsen, G. J., Overbeek, R., Larsen, N., Marsh, T. L., McCaughey, M. J., Maciukenas, M. A., Kuan, W.-M., Macke, J. T., Xing, Y., and Woese, C. R. (1992). The ribosomal database project. Nucleic Acids Res., Suppl. 20, 2199-2200. Pettersson, B., Johansson, K.-E., and Uhlen, M. (1994). Sequence analysis of 16S rRNA from mycoplasmas by direct solid phase DNA sequencing. Appl. Environ. Microbiol. 60, 24562461. Razin, S., Gross, M., Wormser, M., Pollack, Y., and Glaser, G. (1984). Detection of mycoplasmas infecting cell cultures by DNA hybridization. In Vitro 20, 404-408. Sambrook, J., Fritsch, E. P., and Maniatis, T. (1989). "Molecular Cloning: A Laboratory Manual," 2nd ed., Chapter 11. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. Stanbridge, E. J. (1976). A reevaluation of the role of mycoplasmas in human disease. Annu. Rev. Microbiol. 30, 169-187. Van Kuppeveld, F. J. M., van der Logt, J. T. M., Angulo, A. F., van Zoest, M. J., Quint, W. G. V., Niesters, H. G. M., Galama, J. M. D., and Melchers, W. J. G. (1992). Genus- and speciesspecific identification of mycoplasmas by 16S rRNA amplification. Appl. Environ. Microbiol. 58, 2606-2615. Weisburg, W. G., Tully, J. G., Rose, D. L., Petzel, J. P., Oyaizu, H., Young, D., Mandelco, L., Sechrest, J., Lawrence, T. G., Van Etten, J., Maniloff, J., and Woese, C. R. (1989). A phylogenetic analysis of mycoplasmas: Basis for their classification. J. Bacteriol. 171, 64556467. Woese, C. R., Gutell, R., Gupta, R., and Noller, H. F. (1983). Detailed analysis of the higher-order structure of 16S-like ribosomal ribonucleic acids. Microbiol. Rev. 47, 621-669.
A3 CLONED GENOMIC DNA FRAGMENTS AS PROBES David Yogev and Shmuel Razin
Introduction Cloned chromosomal segments have been used as specific probes for the detection and identification of mollicutes (Razin, 1994). The probes are prepared by shotgun cloning of genomic DNA segments and selection of those segments that recognize relatively stable chromosomal regions discriminatory for the specific mollicute species. These species-specific chromosomal segments are not expected to carry rRN A genes or other highly conserved genes. An early attempt to detect noncultivable phytoplasmas (MLOs) in plants with the rRNA gene probe pMC5 failed, as the probe hybridized also with the rRNA genes of the plant chloroplasts (Nur etal., 1986). Further studies succeeded in cloning phytoplasmaspecific chromosomal segments, capable of detection and identification of a variety of phytoplasmas in their plant and insect hosts (Kirkpatrick et al., 1987; Davis et aL, 1990), advancing considerably the research of noncultivable mollicutes (see Chapters E6, Vol. I, and D l l , this volume). For fast and highly efficient construction of a genomic library to be used for the selection of DNA probes, the bacteriophage XGEM-12 with Xhol half-site arms (Promega, Madison, WI) is highly recommended. It has an extremely low background of false recombinants, prevents insert rearrangements, and, in particular, obviates size fractionation of genomic inserts. The preparation of the mycoplasmal DNA requires only partial digestion by the Mbol or Sau3Al restriction enzymes. 47 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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David Yogev and Shmuel Razin
Materials Bacteriophage XGEM-12 with Xhol half-arm sites (Promega) A commercial in vitro k DNA packaging system (available from a variety of commercial sources) A plasmid vector (for example, pGEM, Promega) Nitrocellulose filters (0.05-|xm pore size) 2x SSC (20X SSC is 0.3 M trisodium citrate dihydrate, 3 M NaCl, pH 7.0) 1% sodium dodecyl sulfate (SDS) Denaturation solution (1.5 M NaCl; 0.5 M NaOH) Neutralization solution (1.5 M NaCl; 0.5 M Tris-HCl, pH 7.0) Whatman 3MM filter paper sheets NZCYM 80-mm plates (Sambrook et aL, 1989) overiaid with 3 ml TB top agar (1% Bacto-tryptone; 0.5 ml NaCl; 0.6% agarose; 10 mM MgS04) Conserved gene probes
Procedure 1. Prepare a partial digest of mycoplasmal DNA with Mbol or Sau3Al to generate DNA fragments ranging from 4 to 10 kb. This can be done by establishing the optimum enzyme concentration to generate the desired size range of fragments. 2. Ligate the genomic restriction fragments to Xhol half-arm sites of the XGEM-12 vector. 3. Pack the recombinant phage DNA using a commercial in vitro X DNA package system (Promega). Phage plaques are generated in Escherichia coli strains LE 392 or KW251 on NZCYM agarose plates. 4. Overlay several plates, containing 2 x 10^ plaque-forming units, with nitrocellulose filters for 3 minutes at room temperature. 5. Remove the filters and place them on a Whatman 3MM paper saturated with 1% SDS for 3 minutes (be sure to mark the filters with a needle for correct orientation). 6. Remove the filters and place them on a Whatman 3MM paper saturated with denaturation solution for 5 minutes at room temperature. 7. Repeat step 6 once with neutralization solution and once with 2x SSC. 8. Place the filters on Whatman 3MM paper and allow to dry at room temperature, and then immobilize the DNA by baking the filters in a vacuum oven for 2 hours at 80°C. 9. Duplicates of nitrocellulose filters containing the denatured phage DNA
A3 DNA Probes
49
should be prepared. One set of filters is then hybridized with a mixture of highly conserved gene probes (rRNA genes, such as pMC5; m/gene, etc.) and one set is hybridized with the entire genomic DNA of the most related mycoplasma species. (The hybridization protocol is described in Chapter E5, Vol. I.) 10. A collection of phages which did not react with the highly conserved gene probes and with the genomic DNAs of the most related species are picked up, replated at low density, hybridized again, and subjected to plaque purification. 11. These plaques can then be screened using several other total genomic DNAs of other mycoplasma species usually present in the same host. A phage(s) exhibiting no reaction with any of the mycoplasma species tested can be regarded as a specific probe. 12. The purified phage DNA can be used directly as a DNA probe or restriction segments of the phage DNA insert can be subcloned into a plasmid vector. 13. Confirm the specificity of the recombinant plasmid to be used as a probe by Southern blot hybridization against restricted genomic DNAs of the related mycoplasma species defined earlier.
Discussion Plasmid vectors have been routinely employed in the selection of DNA probes. The availability of genetically engineered commercial bacteriophage vectors provides an alternative way for selecting DNA probes. An example for such a vector, described in this chapter, is the bacteriophage XGEM (Promega). The cloning strategy for this vector depends on the exclusive specificity with which partially filled-in Xhol \ arms can be ligated with partially filled-in Mbol or S^MSAI-digested genomic DNA. The procedure is very rapid, extremely efficient, and requires only small amounts of genomic DNA. The main advantage of this cloning strategy is that it makes genomic DNA fractionation and phosphorylation unnecessary. Within two rounds of plaque purification, a phage which does not contain highly conserved genes or other common sequences of a related mollicute species can be easily obtained. The fact that most pathogenic mycoplasmas colonize the epithelial linings of the respiratory and urogenital tracts would be expected to introduce into the clinical specimens (sputum, urine, swabs, etc.) a significant number of bacteria belonging to the normal flora, including commensal mycoplasmas. This emphasizes the need for imposing rather strict criteria for determining probe specificity, prior to its approval as a routine diagnostic probe. Checking for probe specificity is particularly important with mollicute species exhibiting a significant degree of shared genomic sequences (tested as described in Chapter E5, Vol. I). Examples of mycoplasmas that share genomic sequences and may be present together in the
50
David Yogev and Shmuel Razin
same clinical specimen are Mycoplasma pneumoniae and M. genitalium (Hyman et ai, 1987), M. gallisepticum and M. synoviae (Hyman et al., 1989), M. hyopneumoniae emd M. flocculare (Stemke, 1989; Abiven etal., 1992). In such cases, selection of cloned DNA segments specific to one of the pair should include hybridization of the clones with labeled DNA of the other mycoplasma to rule out cross-reacting clones (Hyman et al., 1987). Cross-reactivity could be overcome by increasing hybridization stringency (Stemke, 1989) and/or by decreasing the size of the DNA segment used as the probe, abating in this way the chances for the occurrence of shared genomic sequences, but at the same time decreasing the sensitivity of the test (Dedieu et ai, 1992). Sensitivity of mycoplasma detection by cloned DNA probes depends on the material tested, hybridization conditions, and on the size and mode of labeling of the probe. When isolated mycoplasmal DNA is tested by dot-blot hybridization, the mycoplasmal probes developed so far could detect down to 0.1-1 ng DNA, corresponding to about 10"^ to 10^ colony-forming units (Hyman et al, 1987, 1989; Stemke, 1989; Abiven et al, 1992; Dedieu et al, 1992; Zhao and Yamamoto, 1993; Razin, 1994). Sensitivity can be increased to some extent by increasing the size of the segment used as a probe or by a composite probe consisting of several species-specific chromosomal segments (Hyman et al., 1987). This may, however, endanger probe specificity. The nonradioactive labeling of probes may decrease sensitivity somewhat compared to that of the same probes labeled by ^2? (Dedieu et. al, 1992). The most crucial parameter of probe effectiveness concerns its ability to detect the specific mycoplasma in clinical samples. The presence of mucus and host cellular debris in the tested specimen requires pretreatment of samples, leading usually to decreased sensitivity to levels significantly lower than those obtained with purified mycoplasmal DNA or with washed organisms. Treatment of tracheobronchial washings of piglets infected by M. hyopneumoniae with 0.5% SDS and proteinase K or testing the DNA extracted from lung homogenates suffered from low levels of detection compared to a diagnosis based on culture, serology, and pathology (Ahrens and Friis, 1991; Abiven et al, 1992). On the other hand, testing of M. capricolum in milk of infected goats required only treatment with NaOH to lyse the mycoplasmas and denature the DNA, retaining the probe sensitivity level (Dedieu et al, 1992). On the whole, the chances of obtaining positive DNA probe tests are better in acute infections, when the number of the mycoplasmas in the specimen is high. In this case, by providing earlier diagnosis, the major advantage of the DNA probe over culture and serology becomes evident (Hyman et al, 1989; Levisohn et al, 1989). Although the value of cloned chromosomal segments as diagnostic probes appears to have decreased with the introduction of the much more sensitive polymerase chain reaction (PCR)-based systems, the cloned segments have retained their value by serving as a basis for establishing target sequences for PCR.
A3 DNA Probes
51
Partial or complete sequencing of a species-specific chromosomal segment enables the selection of an appropriate target sequence and construction of primers for its amplification (Harasawa et al., 1991; Brogan et al, 1992).
References Abiven, P., Blanchard, B., Saillard, C , Kobisch, M., and Bove, J. M. (1992). A specific DNA probe for detecting Mycoplasma hyopneumoniae in experimentally infected piglets. Mol. Cell. Probes 6, 423-429. Ahrens, P., and Friis, N. F. (1991). Identification of Mycoplasma hyopneumoniae with a DNA probe. Lett. Appl. Microbiol. 12, 249-253. Brogan, J. M., Acciai, J., Gallia, G. L., McCleskey, F. K., and Del Vecchio, V. G. (1992). Development of a DNA probe for Ureaplasma urealyticum. Mol. Cell. Probes 6, 411-416. Davis, R. E., Lee, I.-M., Douglas, S. M., and Dally, E. L. (1990). Molecular cloning and detection of chromosomal and extrachromosomal DNA of the mycoplasmalike organism associated with little leaf disease in periwinkle (Catharanthus roseus). Phytopathology 80, 789-793. Dedieu, L., Breard, A., and Lefevre, P. C. (1992). Development of a species-specific DNA probe fox Mycoplasma capricolum. Vet. Microbiol. 32, 189-197. Harasawa, R., Koshimizu, K., Takeda, O., Uemori, T., Asada, K., and Kato, I. (1991). Detection of Mycoplasma hyopneumoniae DNA by the polymerase chain reaction. Mol. Cell. Probes 5, 103-109. Hyman, H. C., Yogev, D., and Razin, S. (1987). DNA probes for detection and identification of Mycoplasma pneumoniae and Mycoplasma genitalium. J. Clin. Microbiol. 25, 726-728. Hyman, H. C., Levisohn, S., Yogev, D., and Razin, S. (1989). DNA probes for Mycoplasma gallisepticum and Mycoplasma synoviae: Applications in experimentally infected chickens. Vet. Microbiol. 20, 323-337. Kirkpatrick, B. C , Senger, D. C., Morris, T. J., andPurcell, A. H. (1987). Cloning and detection of DNA from a nonculturable plant pathogenic mycoplasma-like organism. Science 238, 197200. Levisohn, S., Hyman, H., Perelman, D., and Razin, S. (1989). The use of a specific DNA probe for detection of Mycoplasma gallisepticum in field outbreaks. Avian Pathol. 18, 535-541. Nur, I., Bove, J. M., Saillard, C., Rottem, S., Whitcomb, R. M., and Razin, S. (1986). DNA probes in detection of spiroplasmas and mycoplasma-like organisms in plants and insects. FEMS Microbiol. Lett. 35, 157-162. Razin, S. (1994). DNA probes and PCR in diagnosis of mycoplasma infections. Mol. Cell. Probes 8, 497-511. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). "Molecular Cloning: A Laboratory Manual," 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Stemke, G. W. (1989). A gene probe to detect Mycoplasma hyopneumoniae, the etiological agent of enzootic porcine pneumonia. Mol. Cell. Probes 3, 225-232. Zhao, S., and Yamamoto, R. (1993). Species-specific recombinant DNA probes for Mycoplasma meleagridis. Vet. Microbiol. 35, 179-185.
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A4 PCR: SELECTION OF TARGET SEQUENCES Remi Kovacic^ Odile Grau^ and Alain Blanchard
Target Selection The choice of the DNA target sequences to be amplified depends on the objective of the study. For detection of mycoplasmas in cell cultures, biological fluids, or tissues, the target sequence has to be chosen from sequences that are highly conserved among the strains of the species under study. It is possible to widen the range of species being detected by choosing a target that is conserved among a group of microorganisms. Using conserved sequences of 16S rRNA, it has even been possible to amplify DNA from bacteria that are not cultivable; this approach has been used for the characterization of mycoplasma-like organisms (Schneider et al., 1993, see Chapter E6 in Vol. I and Chapter D l l in this volume). Polymerase chain reaction (PCR)-based assays are not restricted to the detection of a particular species or a group of species since they can be used to detect any genetic character. For example, such an approach has been used for the specific detection of tetracycline resistance in urogenital mycoplasmas; this was feasible because the only genetic determinant for this antibiotic resistance in mycoplasmas is associated with the presence of gene tetM (Blanchard et al., 1992). Nature of Target Sequence The sensitivity of the PCR amplification assay can be improved if more than one target sequence is present in the genome. Therefore, for optimal detection of 53 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. 112
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Remi Kovacic et al.
mycoplasmas, repetitive sequences are the ideal choice. In Mycoplasma fermentans, the presence of a 1405 bp genetic element, which has unique characteristics strongly resembling bacterial insertion sequence-like (IS) elements, has been described. There are from 5 to more than 10 copies of this IS element in the genome of M. fermentans. The sensitivity of the PCR assay based on the amplification of a 206-bp DNA fragment within this IS allows the amplification of 1 fg of genomic DNA which corresponds approximately to a single genome of this microorganism (Wang et al, 1992). With the exception of Mesoplasma lactucae, which has three sets of rRNA genes, the other species have one, or more often two, sets of these genes compared to seven for Escherichia coli. For more than 50 mycoplasma species, the 16S rRNA sequence is available in sequence databases. Some regions of 16S rRNA genes are highly conserved whereas others are variable among species. In addition, there is a high conservation of this gene within a single bacterial species. Therefore, after aligning these sequences, it is usually possible to choose a region within variable domains of the 16S rRNA gene that allows for the specific detection of a particular species. To the contrary, PCR primers chosen in less variable regions will broaden the specificity and allow the detection of more than one mycoplasmal species; this approach has been used by various investigators to develop PCR assays for the detection of mycoplasmal contamination of cell cultures (see Chapter F4, this volume). Problems with this strategy can occur when the sequences from closely related mycoplasmas are not available in databases. This is the case forM. genitalium, which renders difficult the choice of a target within 16S rRNA for the specific detection of the closely related M. pneumoniae. In addition, since there are about 10,000 rRNA molecules per cell, it would be possible in theory to increase the PCR sensitivity by amplifying these targets once cDNA has been made. A protocol using this approach has been published (van Kuppeveld et al., 1992). However, this method has at least one drawback since it requires an additional step which consists of the reverse transcription of rRNAs. Length of Amplified Fragments
The target length can vary from 100 bp to several kbp and the choice of a defined size depends on several parameters: goal of the study, amplification protocol, or need for inactivation of the amplified products. When starting with material that has been treated using protocols that alter the integrity of the DNA, such as formalin fixation, small targets are amplified with more efficiency. For mycoplasmal detection, a length of the target sequence between 100 and 400 bp is sufficient, but for studying the polymorphism among several strains of
A4 PCR: Selection of Target Sequences
55
a given mycoplasma species by restriction analysis or sequence comparison, it may be useful to amplify a larger DNA fragment. However, it should be kept in mind that for targets over 1 kbp, the polymerase activity is more susceptible to low levels of inhibitors, and because of the lack of 5'-^3'-exonuclease activity (proofreading) of the Taq DNA polymerase, these long fragments are more subject to elongation errors. The latter can be minimized with the use of a thermophilic DNA polymerase with proofreading activity such as the UlTma DNA polymerase (Perkin-Elmer, Norwalk, CT). Another factor that can influence the choice of the target length can be the need to inactivate amplified products to avoid false-positive results. Indeed, inactivation using UV irradiation or photochemical treatment by psoralen is more efficient on DNA fragments with a length over 250 bp. Genetic Stability of Target Sequences
Some mycoplasma genes can vary significantly between two strains of the same species. These genes are not a suitable target when DNA amplification from different strains of the same species is desirable. Therefore, it is recommended to choose a DNA sequence that has been shown to be stable among various strains or, at the very least, to have been verified experimentally. When possible, sequences with no known function and chosen randomly should be avoided. Within an open reading frame (ORF), it is preferable to select primers with their 3' end corresponding to the first or the second base of a codon. Indeed, because the pressure of selection is usually directed at the amino acid level and because of the degeneracy of the genetic code, variability is usually more prevalent at the third base of the codons than on other bases. For the same reason, the 3' end of the PCR primer should correspond, when possible, to amino acids with a limited number of possible codons (for example, avoid leucine, arginine, or serine, each of which have six possible codons).
Selection of Primers
Specificity
When choosing PCR primers for the detection of a mycoplasma, its natural habitat should be taken into account. DNA extracted from a sample to be analyzed is likely to contain DNA from the host and also from other microbial agents which inhabit the same site. A method that takes into account the host and
56
M. M. M. M. M. M. U. S. M. A. C. As L.
pi rum gallisepcicum pneumoniae muris lowae penetrans urealycicum ciiri hominis laidlawii innocuum . anaerobium catenaforme
Remi Kovacic e( al.
T G G C G G C A T G C C T A A
JTAGCAATACATT
r
. TG .. G . . G
T. .
C . A ¥ ^'^Olr G C T T G C A C C C . A " '}'Qf A C T T G T G C T T A G -^ .' / ; s ^,t, > ^ V'O C C T T T T . G G . T . A ' "•' \V,* >>' ^ «|C A T C T T C G G A T G C "^i?f5TrTT..GG.AGC. 'TGCGCCN.G. AC . A .v^,^^ w^x v j J C A T T A G G C A C ..•^i.l*.^..<..t«3j^..0^ikS?S^ C T T . G G A G . C C A
Fig. 1 . Selection of a PCR primer with the use of the software PC-Rare. Alignment of M. pirum 16S rDNA (nucleotides 37 to 84) with 16S rDNA of other mycoplasmas and gram-positive microorganisms. The area in gray represents the position of the positive primer for amplification of M. pirum (MYCPIRP) and the corresponding sequences in other microorganisms. The upper graph represents the frequency curve of all octamers of this sequence in the human genome, as determined using the software PC-Rare. The similarity observed between the sequences of M. pirum and M. gallisepticum has no consequence on the specificity of the PCR assay because discrimination of the amplification was obtained with the negative primer.
optimizes the choice of PCR primers with a priori good specificity has been described. This method is illustrated here with the choice of humans as the host (Fig. 1). Using computer analysis, Griffais et al. (1991) determined from among the 4^ possible octamers (65,636) the most and the least frequent sequences from the human genome occurring in the databases, and they selected the 200 most and least frequent. They demonstrated experimentally that this distribution frequency could be extended to the entire human genome. Using primers with 3' ends that correspond to rare sequences in the human genome, specific amplification occurs even in the presence of an excess of human DNA. To the contrary, with PCR primers with 3' end sequences that are frequent in the human genome, a high degree of unspecific amplifications occurred. The PC-Rare software (Eurogentec, Seraing, Belgium) takes into account the K-tuple frequency in a defined environment (human, rodent, Drosophila, Saccharomyces, or E. coli) and determines, in that context, PCR primers whose 3' end octamers are rare, thus having a low probability of nonspecific amplification.
A4 PCR: Selection of Target Sequences
57
Using this program, specific primers have been determined for several molUcute species which can be used in human context without giving unspecific products. Figure 1 shows the frequency curve obtained using this software for the choice of a positive primer for the ampUfication of M. pirum in human material (Grau et al, 1993). Sequences of 16S rDNA obtained from GenBank were aligned using the PileUp software from the GCG package (Genetics Computer Group Inc., Madison, WI), and the alignment shown corresponds to a region with high interspecies variation. The 16S rDNA sequences were chosen among the mycoplasmas phylogetically close to M. pirum, from one of the members of each mollicute phylogenetic group (Weisburg et al., 1989), and from gram-positive bacteria phylogenetically related to mycoplasmas. The specificity and the sensitivity of the PCR assay based on primers chosen using this approach were experimentally verified and, as expected, there was no decrease of sensitivity even in the presence of an excess of human DNA.
Length
Usually the length of an oligonucleotide primer used for PCR varies between 15 and 30 bases. Shorter primers suffer from low specificity, whereas longer ones do not necessarily add specificity, are more expensive to synthesize, and demand an increased annealing time. Primer length is also important when mismatches are added to introduce 5' end sequences corresponding to restriction sites or other modifications. The length of an oligonucleotide primer is largely determined by sequence conservation, size, and composition of the target sequence. If the chosen target is within a sequence highly conserved among mycoplasmas (16S rDNA, for example), the choice of the target length is not of utmost importance. On the other hand, if the chosen amplification sequence is within a very polymorphic region, then the length of the primers is limited by the length of the region showing the polymorphism. Base Composition
DNA melting temperature (T^) for an oligonucleotide bound to its target is the temperature at which 50% of the strands have become separated in solution. T^ depends on several parameters: concentrations of primers and target, primer sequence composition, and PCR mixture composition. Short oligonucleotides, with the same base composition, have lower T^ than longer primers, but this effect is less important than base composition when the length approaches 20 to 25 nucleotides. One of the two formulas used for estimating of T^ is T^ = 81.5°C + 16.6 log[K+] + 0.41(G + C%) - 675/L,
(1)
58
Remi Kovacic et al.
where K+ is the concentration of monovalent salts, G + C% is the G + C% of the primer, and L is the number of bases of the oligonucleotide. The second formula is simplified and only takes into account the base composition of the primer and should be restricted to 15-24 oligomers: T^ = 2°C X (No. of A and T) + 4°C x (No. of C and G).
(2)
However, the determination of T^^ cannot substitute for experimentally testing the optimal annealing temperature of a defined set of primers. This is usually determined by performing a set of experiments with increasing annealing temperatures (by increments of 2°C). The highest temperature showing the optimal sensitivity is the one chosen. It should be mentioned that this temperature is only defined for the particular thermal cycler being used since the way temperatures are expressed from one instrument to another may vary. The mollicutes have an especially low guanine plus cytosine (G + C) content of DNA, ranging from 21 to 43 G 4- C%, with a mean around 30%. The PCR primer specificity can be increased by choosing oligonucleotides having a G + C% which slightly exceeds these values, thus decreasing the probability of crossamplification with other related mycoplasmas. Long stretches of the same nucleotide should be avoided because it may decrease target specificity. The distribution of the four nucleotides must not be too heterogenous. Special attention at the 3' ends, the portion recognized by the polymerases, should be paid in choosing the primers; a high proportion of G + C residues may lead to lower target specificity in providing stable base pairing with nontarget sequences, via (G+C)rich regions. The terminal 3' nucleotide of primers is also critical for good specificity. A mismatch with a 3' thymidine residue is much more tolerated than others (C:C, G:A, or A:A) which could greatly decrease the specificity of the PCR. For the optimization of the choice of the PCR primers, it is recommended to avoid oligonucleotides with complementary structures. These structures favor the self-annealing of primers leading to hairpin loop formation. Specifically, if they are located in the primers 3' end, these structures will reduce the PCR efficiency by lowering the effective primers concentration. The reaction will also be affected if the sequences of the two oligonucleotides are partially complementary, leading to "primer-dimer" formation; this phenomenon is particularly annoying if complementarity is located in the 3' ends of the primers. In order to minimize these problems, three rules must be followed: (i) avoid stable interprimer matching by reducing the complementary contiguous bases number; (ii) avoid even single-nucleotide complementarity at the 3' end between the two primers; and (iii) limit the primers' stable internal hairpin loops formation (Table I). Several commercially available computer programs are able to predict these artifacts and assist with choosing optimal primers (Osborne, 1992; Rychlik and Rhoades, 1989). It also possible to incorporate modified bases that will be used in the detection
A4 PCR: Selection of Target Sequences
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TABLE I GENERAL RECOMMENDATIONS FOR SELECTION OF PCR PRIMERS FOR SPECIFIC DETECTION OF MYCOPLASMAS
Length: 18-25 nucleotides Base composition: G+C% around 50% No long G+C stretches 3' extremity tolerated mismatches T:(G or C), A:C, or C : A » A : A » > C : C , G:A, or A:G No internal complementary sequences Mismatches located at the 5' end
system. For example, 5-bromo-dUTP, digoxigenin-dUTP, or biotin-dUTP can be substituted for TTP. Concentration of PCR Primers Oligonucleotides are often obtained in bulk solutions from the manufacturer. There are several formulas for the calculation of concentration for a synthetic oligomer. Two of the most practical ones are Concentration (M) = OD260 nm ^ D/E^
(3)
or, simplified without taking into account the difference between molar extinction coefficients of the different nucleotides. Concentration
(|JLM)
= OD260 nm/MW x 33 x D x 1000,
(4)
where MW (Da) is the oligonucleotide length x 330; OD260nm is the optical density of the DNA solution at 260 nm; D is the dilution used for OD reading (a 10-|xl aliquot in 1.0 ml would give a D value of 100); and E^ (molar extinction coefficient) is a (16,000) + b (12,000) + c (7000) + d (9600);
(5)
a, b, c and d represent, respectively, the number of A, G, C, and T in the oligomer (this calculation assumes that the oligomer has no secondary structure). Primer Degeneracy In some cases it may be necessary to modify the 5' end of the oligonucleotides for several purposes: (i) addition at the 5' end of a restriction site sequence, usually for cloning purposes. In this case, it is recommended to add two to four extra nucleotides at the 5' end of the primers since restriction enzymes may have difficulty in cleaving at a site located at the far 5' end. If different restriction sites are added to each of the PCR primers, it is possible to clone the amplified DNA
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Remi Kovacic et al.
fragment with a known orientation, (ii) Degenerate primers in the 3' end can be used to amplify related sequences. Purity It is now possible to produce synthetic oligonucleotides with reasonable quality and good yield. It must be taken into account that the best synthetic primers are contaminated with "failure sequences" and modified chains, which usually represent around 10% of the crude product. These undesirable products result from incomplete coupling, leading to truncated oligomers, and chemical side reactions, like depurination, occurring during the synthetic process. These phenomena may reduce signal intensity or permit amplification of unexpected targets. Altered oligonucleotides can be removed through a re versed-phase highperformance liquid chromatography. It is advisable to check the homogeneity of the product on a poly aery lamide gel.
References Blanchard, A., Dybvig, K., Crabb, D. M., and Cassell, G. H. (1992). Rapid detection of tetM in Mycoplasma hominis and Ureaplasma urealyticum by PCR: tetM confers resistance to tetracycline but not necessarily to doxycycline. FEMS Microbiol. Lett. 95, 277-282. Grau, O., Kovacic, R., Griffais, R., and Montagnier, L. (1993). Development of a selective and sensitive polymerase chain reaction for the detection oi Mycoplasma pirum. FEMS Microbiol. Lett. 106, 327-334. Griffais, R., Andre, P. M., and Thibon, M. (1991). K-tuple frequency in the human genome and polymerase chain reaction. Nucleic Acids Res. 19, 3887-3891. Osborne, B . I . (1992). Hyper PCR: A Macintosh HyperCard program for determination of optimal PCR annealing temperature. Comput. Appl. Biosci. 8, 83. Rychlik, W., and Rhoades, R. E. (1989). A computer program for choosing optimal oligonucleotides for filter hybridization, sequencing and in vitro amplification of DNA. Nucleic Acids Res. 17, 8543-8551. Schneider, B., Ahrens, U., Kirkpatrick, B. C , and Seemiiller, E. (1993). Classification of plantpathogenic mycoplasma-like organisms using restriction-site analysis of PCR-amplified 16S rDNA. J. Gen. Microbiol. 139, 519-527. van Kuppeveld, F. J. M., van der Logt, J. T. M., Angulo, A. F., van Zoest, M. J., Quint, W. G. V., Niesters, H. G. M., Galama, J. M. D., and Melchers, W. J. G. (1992). Genus- and speciesspecific identification of mycoplasmas by 16S rRNA amplification. Appl. Environ. Microbiol. 58, 2606-2615. Wang, R. Y.-H., Hu, W. S., Dawson, M. S., Shih, J. W.-K., and Lo, S.-C. (1992). Selective detection of Mycoplasma fermentans by polymerase chain reaction and by using a nucleotide sequence within the insertion sequence-like element. J. Clin. Microbiol. 30, 245-248. Weisburg, W. G., Tully, J. G., Rose, D. L., Petzel, J. P., Oyaizu, H., Yang, D., Mandelco, L., Sechrest, J., Laurence, T. G., van Etten, J., Maniloff, J., and Woese, C. R. (1989). A phylogenetic analysis of the mycoplasmas: Basis of their classification. J. Bacteriol. 171, 6455-6467.
A5 PCR: PREPARATION OF DNA FROM CLINICAL SPECIMENS Bertille de Barbeyrac^ Christiane Bebear^ and David Taylor-Robinson
Introduction Organisms in culture and clinical specimens have to be treated before amplification. Various treatments can be used, according to the organisms to be detected and the kind of specimens in which they occur. Since mycoplasmas do not possess any bacterial cell wall material, DNA may be made available for the polymerase chain reaction (PCR) by simple thermal shock treatment. However, with certain specimens, such as blood or urine, that contain Taq DNA polymerase inhibitors, DNA extraction is necessary. All the procedures must be undertaken in a way that avoids possible carryover of DNA from other samples or laboratory equipment. Three kinds of procedures are described in this chapter, from one that involves little preparation to DNA extraction, according to the samples.
Materials Water bath Centrifuge Lysis buffer: 1 mM EDTA, pH 8.0; 10 mM Tris-HCl, pH 8.0; 0.1% Triton X-100; 200 |JLg/ml proteinase K 61 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Procedure Boiling
Simple boiling can be used for specimens, such as medium in which organisms have been grown. After heating at 95°C for 10 minutes, a 10-|JL1 aliquot can be used directly in the PCR assay. Lysis SPECIMENS
1. Swabs (throat, cervical, urethral, or other) are collected in 2SP medium [0.2 M sucrose in 0.02 M phosphate buffer (pH 7.2) containing 5% fetal calf serum] or in mycoplasmal culture medium. 2. Cellular material from fluids, such as bronchoalveolar lavage (BAL) and cerebrospinal fluid (CSF), is recovered by centrifugation (3500 ^ for 15 minutes). The deposit is resuspended in 1 ml of supernatant fluid. 3. Small pieces of tissue (for example, 1 x 10 mm) are taken at biopsy. The tissues in phosphate-buffered saline (PBS) are disintegrated by vortexing with glass beads and then centrifuged as described earlier. METHOD
Aliquots of 200 |xl of these specimens are centrifuged at 13,000 g for 30 minutes. The deposits are resuspended in 100 jxl of lysis buffer and the mixtures are incubated at 55°C for 90 minutes. After heating at 95°C for 10 minutes, lO-jxl aliquots are subjected to PCR amplification (de Barbeyrac et al., 1993).
Nucleic Acid
Extraction
SPECIMENS
1. The urine (20 ml) deposit is obtained by centrifugation as described for BAL or CSF. 2. Peripheral blood mononuclear cells (PBMC) are obtained by FicollHypaque centrifugation from whole blood collected in a tube containing EDTA. 3. Large pieces of tissue taken at biopsy. The tissue is cut into smaller pieces which are disintegrated and processed as described earlier. METHOD
The deposits of cells from urine or a disintegrated biopsy specimen or from deposits of PBMC are suspended in 500 |xl of lysis buffer and are purified by
A5 PCR: Preparation of DNA
63
phenol and phenol/chloroform extraction, followed by precipitation with 3 M sodium acetate and 95% ethanol. After lysis, an equal volume of Tris-saturated phenol is added, and the mixture is vortexed gently for 1 minute until a fine emulsion is created. The organic phase is separated by centrifuging at low speed for 5 minutes and the upper aqueous phase is transferred to a new clean tube. An equal volume of phenol/chloroform/isoamyl alcohol is added and mixed in the following ratio according to volume (25:24:1). After mixing, the solution is centrifuged as described earlier and a one-tenth volume of 3 M sodium acetate and 2 volumes of cold 95% ethanol are added. After thorough mixing, the DNA is allowed to precipitate at — 20°C for at least 1 hour. The DNA is recovered by centrifugation at 13,000 g for 30 minutes and the deposit is resuspended in 100 |JL1 of TE buffer. A lO-jxl aliquot of the DNA preparation is used in the PCR assay.
Discussion Boiling is the simplest way of making DNA available but it does not eliminate inhibitory factors of Tag DNA polymerase and is not recommended for clinical samples. For these, the most simple method involves the use of nonionic detergents and proteinase K to solubilize DNA from cells in PCR buffer. Tween 20, nonidet P-40 (NP-40), or Triton X-100 are compatible with Taq DNA polymerase. Jensen et al. (1991) and Katseni et al. (1993) used NP-40 and Tween 20 in their lysis buffer. Unfortunately, the commonly used detergent, sodium dodecyl sulfate, cannot be used. Proteinase K is a very good protease for digesting cells to release DNA or RNA into a form readily accessible to the polymerase. It has the advantage of being relatively stable in the mid-temperature range (50° to 60°C), yet can be thermally inactivated easily at 95°C. The inactivation of proteinase K is a very important step, and Williamson et al. (1992) have shown that the inclusion of 0.01% (v/v) NP-40 in the buffer used for proteinase K digestion promotes full inactivation of proteinase K during boiling. DNA isolated by the detergent/protease method is as suitable as purified material for the PCR. However, urine and blood possess considerable inhibitory activity and the nucleic acids must be purified. The drawback of DNA purification through phenol/chloroform and the final ethanol precipitation step is the risk of a significant loss of DNA. The chelating resin Chelex 100 may provide a better amplifiable yield. A one-step microbial DNA extraction method using Chelex 100, which is suitable for PCR amplification, has been described by de Lamballerie et al. (1992). This method could be an alternative to bacterial lysis followed by phenol/chloroform extraction and ethanol precipitation of nucleic acids, but has not yet been applied to the detection of mycoplasmas in clinical samples. Heparin appears to be a potent inhibitor of gene amplification reactions when
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Bertille de Barbeyrac et al.
nucleic acids are extracted from cells or plasma collected in heparinized tubes. EDTA, which lacks inhibitory activity, is the most appropriate anticoagulant for blood samples when PCR amplification is anticipated (Holodniy et al., 1991).
References de Barbeyrac, B., Bemet-Poggi, C , Febrer, F., Renaudin, H., Dupon, M., and Bebear, C. (1993). Detection of Mycoplasma pneumoniae and Mycoplasma genitalium in clinical samples by polymerase chain reaction. Clin. Infect. Dis. 17, (Suppl. 1), 83-89. de Lamballerie, X., Zandotti, C , Vignoli, C , Bollet, C , and de Micco, P. (1992). A one-step microbial DNA extraction method using "Chelex 100" suitable for gene amplification. Res. Microbiol. 143, 785-790. Holodniy, M., Kim, S., Katzenstein, D., Konrad, M., Groves, E., and Merigan, T. C. (1991). Inhibition of human immunodeficiency virus gene amplification by heparin. J. Clin. Microbiol. 29, 676-679. Jensen, S. J., Uldum, S. A., Sondergard-Andersen, J., Vuust, J., and Lind, K. (1991). Polymerase chain reaction for detection of Mycoplasma genitalium in clinical samples. J. Clin. Microbiol. 29, 46-50. Katseni, V. L., Gilroy, C. B., Ryait, B. K., Aryoshi, K., Bieniasz, P. D., Weber, J. N., and TaylorRobinson, D. (1993). Mycoplasma fermentans in individuals seropositive and seronegative for HIV-1. Lancet 341, 271-273. Williamson, J., Marmion, B. P., Worswick, D. A., Kok, T. W., Tannock, G., Herd, R., and Harris, R. J. (1992). Laboratory diagnosis oi Mycoplasma pneumoniae infection. 4. Antigen capture and PCR-gene amplification for detection of the mycoplasma: Problems of clinical correlation. Epidemiol. Infect. 109, 519-537.
A6 PCR: AMPLIFICATION AND IDENTIFICATION OF PRODUCTS Bertille de Barbeyrac and Christiane Bebear
Amplification Many protocols are described in the literature for the detection of human, animal, and plant mollicutes by polymerase chain reaction (PCR). The principle is always the same, but the experimental conditions differ. The PCR parameters to be optimized include the composition of the mixture (dNTP, Taq DNA polymerase, assay buffer), the volume used, and the amplification cycle (temperature of the different steps, number of cycles). The protocol described in this chapter has been used to detect Mycoplasma pneumoniae and M. genitalium in clinical specimens (de Barbeyrac et al, 1993). Alternative procedures are considered in the discussion section. Materials Thermocycler Three separate rooms (two for pre-PCR manipulations, sample preparation, and mixture preparation, and one for post-PCR manipulations and identification of amplified products) Positive displacement pipettes or pipettes with plugged tips Procedure The following standard PCR protocol can be used for the amplification of most DNA target sequences: 65 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Bertille de Barbeyrac and Christiane Bebear
1. Set up a 50-|JL1 reaction mixture in a 0.5-ml microfuge tube. The mixture consists of 1 |xM of each primer (1 jxl from a 50 jxM stock solution); 200 [xM of each dNTP (1 |xl from a 10 mM stock solution); 1.5 mM MgCl2 (3 JJLI from a 25 mM stock solution); i x assay buffer (5 |xl from a lOx assay buffer without MgCl2); and 1 unit of Tag DNA polymerase. Mix and overlay with two drops of mineral oil. In another room, add 10 jxl of template DNA to the tube (clinical samples or positive control). 2. After 5 minutes of denaturation at 95°C, perform 35 cycles using the following temperature profile: denaturation 95°C, 1 minute; primer annealing 55°C, 1 minute; and primer extension 72°C, 1 minute. Cycling should be concluded with a final extension at 72°C for 5 to 10 minutes. 3. Controls: It is imperative that diagnostic assays include controls to verify results. A negative control (negative sample, reaction mixture without template) and a positive control (culture of a mycoplasma strain or purified DNA) must be included in each PCR experiment.
Discussion
It is highly advantageous to optimize the PCR conditions for a given application. The following points should be considered. CONCENTRATION OF DIFFERENT COMPONENTS /. Enzyme
Concentration
A recommended concentration range for Tag DNA polymerase is between 1 and 2.5 units per tube. It is recommended to test enzyme concentrations ranging from 0.5 to 5 units per reaction. It is essential for diagnostic assays using different patient samples to use an excess of polymerase because of the potential inhibitory activity of the sample contents. 2. Deoxynudeotide
Triphosphates (dNTP)
The optimal dNTP concentration depends on the length of the amplified product and MgCl2 and primer concentrations. It is recommended to test a concentration range for dNTP between 20 and 200 |JLM each. Use a 10 mM stock solution, equimolar for each of the four dNTPs. 3. Magnesium
Concentration
The MgCl2 concentration may affect several parameters such as primer annealing, strand dissociation temperature of both template and PCR products,
A6 PCR Procedure for Mycoplasma Detection
67
product specificity, formation of primer-dimer artifacts, and enzyme activity and fidelity. The optimum MgCl2 concentration may range from 0.5 to 5 mM. 4. Primer
Concentration
Generally, suitable primer concentrations are between 0.1 and 1 [LM. In many cases, the optimum primer concentration is related to the length of the amplified fragment. For optimized assays, residual nonextended primers should be visible on the gel after PCR, whether there is an amplified product or not. In optimizing the conditions for multiplex PCR, the relative concentration of the pairs of primers is the most important variable, as described by Cadieux et al. (1993) for the detection of M. pneumoniae and M. genitalium. 5. Assay Buffer
The recommended buffer for PCR is 10 to 50 mM Tris-HCl (between pH 8.3 and 9) and 50 mM KCl. Gelatin or bovine serum albumin and nonionic detergents such as Tween 20 or Triton X-100 are included to stabilize the enzyme. Generally, the assay buffer is supplied with the Tag DNA polymerase by the manufacturer. CYCLE CONDITIONS, TIME, A N D TEMPERATURE
It is necessary to adapt the cycle conditions, time, and temperature to the tubes and PCR machines that are being used. /. Denaturation
Step
Typical denaturation conditions are 95°C for 30 seconds. However, higher temperatures may be appropriate, especially for (G + C)-rich targets. Denaturation steps that are too high and/or too long lead to an unnecessary loss of enzyme activity. The half-life of Taq DNA polymerase activity at 92.5°, 95°, and 97.5°C is 2 hours, 40 minutes, and 5 minutes, respectively. 2. Primer Annealing and Extension
The temperature and time required for primer annealing depend on the base composition, length, and concentration of the primers and the specificity required. An applicable annealing temperature is 5° to 10°C below the T^ of the primers. Primer extension is traditionally performed at 72°C. Using only two temperatures, a PCR may be run at higher annealing temperatures and should be performed between 55° and 75°C for annealing and extension and at 95°C for denaturation.
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Bertille de Barbeyrac and Christiane Bebear
TARGET
RNA can be used as a target. However, as RNA cannot serve directly as a template for PCR, the successful combination of reverse transcription (RT) and PCR converts RNA into a complementary DNA (cDNA) amenable to PCR. The major advantage of RT-PCR is the sensitivity of the method because of the presence of high copy numbers of RNA molecules in the cell. Tjhie et al. (1994) compared the sensitivity of a direct PCR and a RT-PCR for the detection of M. pneumoniae using either 16S rDNA or 16S rRNA as targets. In this experiment, the RT reaction was performed in a 20-|JL1 volume containing 20 units of RNasin (RNase inhibitor) and 8 units of avian myeloblastosis virus reverse transcriptase; only one-fourth of the RT reaction mixture was used in the PCR. A disadvantage of RT-PCR is the need for RNA purification. Furthermore, all precautions designed to avoid RNA degradation or contamination must be strictly followed. CONTROLS Negative
Controls
Negative controls must be used to verify the absence of contamination pre- or post-PCR. A minute amount of target DNA contaminating the sample or reagents will give rise to false-positive results. All PCR tests must be carried out with strict precautions to prevent contamination (separate rooms, positive displacement pipettes or pipettes with plugged tips). The use of chemical modification of PCR products prevents carryover of amplifiable fragments. The addition of dUTP instead of dTTP during PCR allows the production of a modified DNA. This modified DNA can be digested with the enzyme uracil A^-glycosylase (UNG). If UNG is added to the reaction mixture, before a novel amplification, carried-over fragments are digested, but not the sample DNA. This is the principle of a commercially available carryover prevention kit (Perkin-Elmer). Positive Controls
The nature and quality of the positive control are a crucial point. Falsenegative results can be caused by known or unknown parameters such as a malfunction of the thermal cycler, error in the composition of the reagents, batch-related variability of reagents (primers and Taq polymerase), mishandling during extraction of DNA from the samples, or inhibitors present in the samples tested. A positive culture of the tested mycoplasma suspended in the same reaction mixture as the samples can serve as a positive control. This kind of external positive control cannot detect false-negative results due to inhibitors of
A6 PCR Procedure for Mycoplasma Detection
69
Taq polymerase introduced by the clinical material. To prevent false-negative results due to inhibitors present in the sample tubes, it is possible to use human genes (globin, HLA) as internal positive controls either in a simplex PCR or in multiplex PCR as described by Cadieux et al. (1993). With the latter method, several genomic regions from infectious agents or from a eukaryotic gene can be amplified simultaneously in one tube using a set of different primer pairs. It is important that the human gene PCR amplifies a fragment comparable in length to the fragment amplified in the specific PCR. This kind of control indicates if the tested sample is of good quality. Ursi et al. (1992) have described an internal positive control that can be amplified with the same pair of primers as the target DNA, but giving a different-sized PCR product. The control is a constructed plasmid containing the specific M. pneumoniae amplicon, added to each sample tube. The drawback of this control is the potential competition between the primers on the same target.
Identification of Products The PCR-amplified fragment may be visualized on an electrophoresis gel by ethidium bromide staining. The choice of gel, agarose or acrylamide, depends on the fragment size. Several DNA molecular weight markers suitable for approximate size analysis of PCR products are commercially available. This method is usually not sufficient to verify specificity. After gel analysis, the identity of a fragment is checked by restriction enzyme digestion and/or routine blot transfer of the gel for hybridization analysis. In the protocol taken as an example, the expected 466-bp PCR product is visualized on a 1.2% agarose gel, digested with the Taql restriction enzyme (three fragments are visualized on acrylamide gel), and hybridized with a ^^P-radiolabeled internal probe. Other procedures for detection of the amplified fragment are discussed.
Materials EQUIPMENT
Electrophoresis apparatus Microwave oven Hybridization oven Water bath Nylon membrane
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Bertilie de Barbeyrac and Christiane Bebear
REAGENTS A N D BUFFERS
TAE 50X: 242 g Tris base, 57.1 ml glacial acetic acid, and 100 ml 0.5 M EDTA, pH 8.0; bring to 1 liter with H2O 20X SSC: 175.3 g NaCl and 88.2 g sodium citrate; adjust the pH to 7.0 and the volume to 1 liter with H2O Gel-loading buffer: 50% glycerol in H2O and 0.1% bromophenol blue 26% acrylamide stock solution: 25 g acrylamide and 1 g A^,A^'-methylene bisacrylamide; bring to 100 ml with H2O Agarose (electrophoresis grade)
Procedure ELECTROPHORESIS
Before loading the DNA samples into the wells of the submerged gel, mix the sample (10 |xl) with the gel-loading buffer (2 |xl) that serves to increase the density and color of the sample so that the migration rate can be monitored. Run the gel until the bromophenol blue has migrated to the appropriate distance through the gel. After migration, stain the gel with ethidium bromide (0.5 |xg/ml) for a few minutes at room temperature and then photograph the gel. Agarose Gel
Seal the gel tray and place it on a horizontal surface. Mix the agarose at the desired concentration (w/v) with 1 x electrophoresis buffer and heat in a microwave oven. Cast the gel into a prepared tray. Acrylamide Gel
Position the spacers between the plates, fix the plates with clips, and place them in a vertical position. Mix
26% acrylamide stock solution TAE 50X 10% ammonium persulfate TEMED H.O
13.8 ml 900 [xA 405 M-l 4 5 fjLl
29.85 ml
Cast the gel and wait for polymerization to take place at room temperature.
A6 PCR Procedure for Mycoplasma Detection
71
CONTROL OF SPECIFICITY Restriction
Analysis
Set up an aliquot of PCR product (9 |JL1), mix with an optimal reaction buffer generally supplied by the manufacturer to yield a i x end concentration (1 jxl), and add 5-10 units (1 |JL1) of restriction enzyme (Taql). Incubate the mixture for at least 2 hours at optimal temperature (60°C) and visualize the cleavage products on an ethidium bromide-stained acrylamide gel. A positive control that will allow evaluation of enzyme activity is recommended. This control of specificity is possible only when the amplified product possesses a restriction site in an adequate position. Hybridization
Analysis
The DNA from the agarose gel can be transferred using the capillary transfer method (Sambrook et al, 1989) or more rapidly under vacuum. The choice of blotting and hybridization buffers depends on the nature of the membrane used (see also Chapter E5, Vol. I). The nylon membrane (Gene Screen Plus, Nen Research Products, Du Pont de Nemours, Germany) is hybridized overnight at 55°C in 5 ml (50 (xl/cm^) of 1% sodium dodecyl sulfate (SDS), 10% dextran sulfate, 1 M NaCl containing 5 x 10^ dpm/ml of ^^p-labeled oligonucleotide, and 100 |jLg of sheared carrier DNA, such as salmon sperm DNA, in a hybridization oven. The membrane is washed twice for 5 minutes at room temperature in 2x SSC, twice for 30 minutes at 55°C in 2x SSC, 1% SDS, and then for 30 minutes at room temperature in 0.1 x SSC. After the final rinse, the blot is exposed to X-ray film. The synthetic oligonucleotide is labeled by phosphorylation at the 5' end using the bacteriophage T4 polynucleotide kinase. Set up a reaction mixture as follows oligonucleotide (10 pmol/jxl) lOX buffer T4 polynucleotide kinase [7-32p]ATP (specific activity 5000 Ci/mmol)
H2O
Composition of lOX buffer:
10
|JL1
3 M-1 1 fxl 10 \iX 6 \x\
0.5 M Tris-HCl, pH 7.6, 0.1MMgCl2 0.125 mM dithiothreitol, and 1 mM spermidine
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Bertille de Barbeyrac and Christiane Bebear
Incubate for 30 minutes at 37°C and then for 10 minutes at 65°C to inactivate the T4 kinase. The eUmination of unincorporated radioactivity from the oligonucleotide can be achieved by precipitation with ethanol if the oligonucleotide is more than 18 nucleotides in length or by chromatography through a commercially available column.
Discussion
In routine assays, a direct blot of PCR products (dot or slot blot) may be advantageous. This procedure requires no electrophoresis and no DNA transfer, but the drawback is that the PCR product size cannot be visualized. In the protocol of Williamson et al. (1992), applied to M. pneumoniae, the amplified product was examined by quantitative dot-blot hybridization using a synthetic 32? hairpin probe. The specific activity of this probe is on average 10 times higher than that of 5' end-labeled probes. In the protocol of Luneberg et al. (1993), also applied to the detection of M. pneumoniae, the PCR fragments were generated with one biotinylated primer and one unlabeled primer. After the removal of unincorporated primers (Gene Clean), the amplified fragments were immobilized on a streptavidin-coated microtiter plate and were revealed by subsequent hybridization with a digoxigeninlabeled oligonucleotide. The detection of hybrids was performed using the alkaline phosphatase conjugated to the anti-digoxigenin antibody for which the substrate was 4-methylumbelliferyl phosphate.
References de Barbeyrac, B., Bemet-Poggi, C , Febrer, F., Renaudin, H., Dupon, M., and Bebear, C. (1993). Detection of Mycoplasma pneumoniae and Mycoplasma genitalium in clinical samples by polymerase chain reaction. Clin. Infect. Dis. 17, (Suppl. 1), S83-S89. Cadieux, N., Lebel, P., and Brousseau, R. (1993). Use of a triplex polymerase chain reaction for the detection and differentiation of Mycoplasma pneumoniae and Mycoplasma genitalium in the presence of human DNA. J. Gen. Microbiol. 139, 2431-2437. Luneberg, E., Jensen, J., and Frosch, M. (1993). Detection oiMycoplasma pneumoniae by polymerase chain reaction and non radioactive hybridization in microliter plates. J. Clin. Microbiol. 31, 1088-1094. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989). "Molecular Cloning: A Laboratory Manual," 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Tjhie, J. H. T., van Kuppeveld, F. J. M., Roosendaal, R., Melchers, W. J. G., Gordijn, R., MacLaren, D. M., Walboomers, J. M. M., Meijer, C. J. L. M., and van den Brule, A. J. C. (1994). Direct PCR enables detection of Mycoplasma pneumoniae in patients with respiratory tract infections. J. Clin. Microbiol. 32, 11-16. Ursi, J. P., Ursi, D., leven, M., and Pattyn, S., R. (1992). Utility of an internal control for the polymerase chain reaction. Acta Pathol. Microbiol. Immunol. Scand. 100, 635-639.
A6 PCR Procedure for Mycoplasma Detection
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Williamson J., Marmion, B. P., Worswick, D. A., Kok, T. W., Tannock, G., Herd, R., and Harris, R. J. (1992). Laboratory diagnosis of Mycoplasma pneumoniae infection. 4. Antigen capture and PCR-gene amplification for detection of the mycoplasma: Problems of clinical correlation. Epidemiol Infect. 109, 519-537.
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A7 PCR: APPLICATION OF NESTED PCR TO DETECTION OF MYCOPLASMAS Ryo Harasawa
Introduction One of the most popular modifications of polymerase chain reaction (PCR), known as nested PCR (Mullis and Faloona, 1987), is an in vitro DNA amplification employing nested primer pairs. In a typical protocol for the nested PCR, a first-round PCR is performed with a pair of outer primers. A small amount of the first-round PCR product is transferred to a fresh reaction tube for a second-round PCR by using a pair of inner primers. The nested PCR is currently the most sensitive means of detecting the mycoplasmas in cell cultures or in clinical materials because a single copy of target sequence can be detected (Harasawa et al., 1993a,b). Another advantage of the nested PCR is that the second-round PCR serves to confirm the specificity of the first-round PCR. The nested primer pairs recommended in this chapter consist of sequences of the 16S-23S rRNA intergenic spacer regions (Fig. 1).
Materials For Isolation of DNA from Mycoplasma-Contaminated Biological Materials Biological materials (cell cultures, serums, vaccines, and other biologicals) Sterile water, injection or irrigation grade (Note: Conventional distilled water may occasionally be contaminated with trace amounts of prokaryotic DNA) 75 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Spacer region 16SrRNAgene1 Fl
1 23S rRNA gene
,
.
F2
Rl
R2
Probe Fig. 1. Orientation and location of the nested primer pairs and the internal probe (Harasawa et a/., 1993a).
Sodium dodecyl sulfate (SDS) solution, 10% Sodium acetate, 3 M Mussel glycogen, 20 mg/ml (Boehringer Mannheim GmbH) Ethanol, 75% 2-Propanol, absolute Ice bath Microcentrifuge (e.g., Eppendorf centrifuge) capable of 12,000 g Microtube (e.g., Eppendorf tube, 1.5-ml volume)
For Polymerase Chain Reaction
Tag polymerase Reaction buffer, 10 x (supplied with the enzyme) Magnesium chloride, 25 mM Sterile water, injection or irrigation grade dNTPs solution, 2.5 mM each Primers for the first-round PCR Primer Fl [5'-ACACCATGGGAG(C/T)TGGTAAT-3'], 10 pmol/jjil Primer Rl [5'-CTTC(A/T)TCGACTT(C/T)CAGACCCAAGGCAT-3'], 10 pmol/|JLl Primers for the second-round PCR Primer F2 [5'-GTG(C/G)GG(A/C)TGGATCACCTCCT-3'], 10 pmol/jxl Primer R2 [5'-GCATCCACCA(A/T)A(A/T)AC(C/T)CTT-3'], 10 pmol/|xl Internal probe [5'-GTTCTTTGAAAACTGAAT-3'] Mineral oil (e.g., Sigma St. Louis, MO M3516) DNA thermal cycler (Perkin-Elmer Cetus, Norwalk, CT) Microtube (0.5-ml volume)
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71
For Agarose Gel Electrophoresis Agarose (e.g., NuSieve 3:1) Ethidium bromide solution, 0.4 |JLg/ml Dye solution, 6x (0.25% xylene cyanol, 0.25% bromophenol blue and 40% sucrose in water) Tris-acetate buffer, lOx (400 mM Tris, pH 8.0, 50 mM sodium acetate, 10 mM disodium EDTA) Gel electrophoresis apparatus UV transilluminator Camera (e.g., Polaroid land camera MP4) Filter (e.g., Wratten gelatin filter No. 23A, Kodak Rochester, NY)
Procedure Isolation of DNA DNA extraction is usually not necessary in PCR tests for detecting mycoplasmas in fresh (not frozen) cell cultures. It is, however, recommended to carry out this step on testing clinical materials or biological products which may contain inhibitors of PCR. 1. Transfer 450 |JL1 of the tested biological material into a sterile Eppendorf tube prior to subculturing. [Note: Cell cultures grown for 3-5 days after passage are suitable for direct testing. Cell debris from frozen cell cultures may inhibit PCR. A single dose of lyophilized viral vaccines or other biologicals should be dissolved in 450 (JLI of reconstituent (injection grade water) and subjected to DNA isolation. Fetal bovine serum can be directly subjected to PCR without DNA extraction.] 2. Add 50 |xl of 10% SDS solution into the Eppendorf tube, and mix with the tested material for 10 seconds by hand. 3. Cool the tube in ice for 5 minutes to lyse the mycoplasma cells. 4. Add 250 JULI of 3 M sodium acetate solution, and vortex the tube for 5 seconds. 5. Cool the tube in ice for 30 minutes. 6. Centrifuge the tube at 12,000 g for 10 minutes to precipitate the SDSsodium acetate-cell debris complex. 7. Transfer 600 |JL1 of the supernatant fluid into a fresh Eppendorf tube. 8. Add 1 U | L1 of 20 mg/ml mussel glycogen and 600 |JL1 of absolute 2-propanol. 9. Cool the tube in ice for 30 minutes to precipitate DNA.
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Ryo Harasawa
10. Centrifuge the cooled mixture at 12,000 g for 15 minutes. 11. Discard the supernatant fluid, and wash the pellet three times with 600 |xl of 75% ethanol. (A white pellet can be observed at the bottom of the tube.) 12. The pellet is allowed to air dry to remove ethanol. 13. Dissolve the pellet in 10 JJLI of sterile distilled water. 14. Measure the absorbance of the DNA solution at 260 and 280 nm, if necessary. (Note: Use 1 jxl of DNA solution for this purpose.) 15. Dilute DNA to a final concentration between 50 and 250 ng/jxl so that the DNA may be added to the PCR reaction mixture in a volume of 5 (xl or less.
Nested PCR
1. Mix 4-5 |xl of the DNA solution with 10 pmol of each of the first-round PCR primers in a total volume of 50 |xl containing 5 jxl of 10 x PCR buffer, 1 mM of each dNTP, 4 JJLI of 25 mM MgCl2, and water. (Note: When preparing the PCR mix, it is recommended to make allowances for losses of volume by preparing enough PCR mix for one or two samples more than required.) 2. Heat to 94°C for 5 minutes and briefly centrifuge. 3. Add 1 unit of Tag DNA polymerase, mix gently. 4. Add two drops (ca. 40 |xl) of mineral oil. 5. Perform 30 cycles of denaturation (at 94°C for 30 seconds), annealing (at 55°C for 2 minutes), and primer extension (at 72°C for 2 minutes). The final 72°C incubation is extended for 5 minutes. 6. Take 1 jxl of the first-round PCR product to initiate second-round PCR using primers F2 and R2. The composition of the second-round PCR mix is the same as that of the first-round except for the primers. (Note: Strict precautions should be taken to avoid cross-contamination with aerosolized first-round PCR products.) 7. Heat at 94°C for 30 seconds. 8. Add 1 unit of Taq DNA polymerase. 9. Add two drops (ca. 40 |xl) of mineral oil. 10. Perform 30 cycles of denaturation (at 94°C for 30 seconds), annealing (at 55°C for 2 minutes), and primer extension (at 72°C for 2 minutes). The final 72°C incubation is extended for 5 minutes. 11. When cycling is completed, remove 10 |xl from each tube of the first- and second-round PCR and add 2 |JL1 of dye solution. Electrophorese the entire amount on a 1.0% agarose or a 2.0% NuSieve 3:1 agarose gel for 45 to 50 minutes at 50 V in Tris-acetate buffer until the dye front of the bromophenol blue migrates half the length of the gel. 12. Stain the gel in the ethidium bromide solution for 15 to 30 minutes. (Note: Destain the gel in water for 5 minutes, if necessary.)
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13. Photograph the gel on a UV transilluminator using a Polaroid camera equipped with a Wratten gelatin filter No. 23A.
Discussion The DNA extraction step can be skipped if the sample is expected to be free of any inhibitors of PCR. High-titerd mycoplasma cultures may inhibit the PCR because intracellular mycoplasmal components can inhibit the activity of Taq DNA polymerase. The common mycoplasma species infecting cell cultures can be identified by analyzing the restriction patterns of the second-round PCR products (Harasawa et al, 1993a). Although the nested PCR may increase the risk of cross-contamination with product DNA because of its high sensitivity, it does provide a faster means of detecting low-titer infections and, in addition, it confirms the specificity of the first-round PCR (Harasawa et al, 1993b). To avoid cross-contamination, it is good laboratory practice to perform the first-round and second-round amplifications in two separate rooms with different sets of micropipettes. In conclusion, strict precautions should be taken to avoid contamination.
References Harasawa, R., Mizuasawa, H., Nozawa, K., Nakagawa, T., Asada, K., and Kato, I. (1993a). Detection and tentative identification of dominant mycoplasma species in cell cultures by restriction analysis of the 16S-23S rRNA intergenic spacer regions. Res. Microbiol. 144, 489493. Harasawa, R., Uemori, T., Asada, K., and Kato, I. (1993b). Sensitive detection of mycoplasmas in cell cultures by using two-step polymerase chain reaction. In "Rapid Diagnosis of Mycoplasmas" I. Kahane and A. Adoni, (eds.) pp. 227-232. Plenum, New York. Mullis, K. B., and Faloona, F. (1987). Specific syntheis of DNA in vitro via a polymerase-catalyzed chain reaction. In "Methods in Enzymology" (R. Wu, ed.), Vol. 155, pp. 335-350. Academic Press, Orlando, PL.
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A8 PCR: RANDOM AMPLIFIED POLYMORPHIC DNA FINGERPRINTING Steven ] Geary and Mark H. Forsyth
Introduction The accurate identification of strains of mycoplasmas within established species is essential to the studies of epidemiology, pathogenesis, and taxonomy. Traditional methods of identification include serodiagnosis, protein profile comparisons, Western blots, and DNA probes (Chapters E4 and E5 in Vol. I). Serodiagnosis is simple and fast; however, it is relatively insensitive and inadequate for isolates that do not fall within an existing serogroup. Protein profiles and Western blots require large samples, are time-consuming, and are often unable to distinguish between closely related strains. Specific DNA probes possess the required sensitivity and are relatively rapid but again are unable to distinguish or type newly emerging strains. One of the most promising analytical methods for the detection of DNA sequence diversity among mycoplasmas involves polymerase chain reaction (PCR) amplification using a single primer under conditions of low stringency. The primer stimulates the synthesis of DNA fragments from paired sites (in opposite orientations) in the target genome sufficiently homologous to itself to allow for annealing and extension. This results in strain-specific arrays of DNA fragments that can reproducibly distinguish even closely related strains of a species (Welsh and McClelland, 1990; Williams et ai, 1990). This method is commonly referred to as either random amplified polymorphic DNA (RAPD) or arbitrary primer polymerase chain reaction. It is rapid and sensitive, requires only nanogram quantities of template DNA, and is useful for strain identification within numerous genera (Micheli et al., 1993; 81 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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Akopyanz et al., 1992a,b; Welsh et aL, 1992; Wang et al., 1993; Kersulyte et al., 1992; Geary et aL, 1994).
Materials Thermal cycler Oligonucleotide primers* (short, i.e., 10-mers) AmpliTaq DNA polymerase (Perkin-Elmer Cetus, Norwalk, CT) Nucleotides (dATP, dCTP, dGTP, and dTTP) Electrophoresis apparatus UV illuminator TAE [40 mM Tris (pH 7.6), 20 mM sodium acetate, 2 mM EDTA]
Procedure 1. Set up for a total reaction volume of 50 jxl in a sterile 0.5-|xl microfuge tube. 2. Add 2.5 units of AmpliTaq DNA polymerase (Perkin-Elmer Cetus); 250 |xM each of dATP, dCTP, dOTP, and dTTP; 3 mM MgCl2; 400 ng primer; and 40 ng of template DNA. Overlay with 50 jxl of sterile mineral oil. 3. Place the microfuge tube into the thermal cycler programmed for the appropriate amplification reactions. An example of a typical set of reaction conditions begins at 97°C for 5 minutes. Add Taq polymerase, 4 cycles of 94°C for 5 minutes, 36°C for 5 minutes, and 72°C for 5 minutes followed by 30 cycles of 94°C for 1 minute, 36°C for 1 minute, and 72°C for 1 minute followed by 1 cycle of 72°C for 10 minutes. 4. Thirty-microliter aliquots of amplified DNAs are electrophoresed in 2% (w/v) agarose gels in TAE containing 0.5% ethidium bromide (v/v) at 6 V/cm. One microgram of a 123-bp DNA ladder (GIBCO/BRL) should also be run in each gel as a standard for size determination of DNA fragments. 5. Visualize the DNA with a UV illuminator at 300 nm. 6. Photograph. *Many individual primers may have to be tested to obtain optimum results. The primer should be short and selected with consideration for the G + C ratio of the species to be analyzed. Mycoplasmas in general are A + T-rich, therefore, a primer which is, for example, 70% A + T may result in too many bands compared to one which is only 40% A + T.
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Discussion Accurate and reproducible RAPD fingerprinting patterns require that the mycoplasma be propagated in pure culture prior to DNA isolation. Contaminating DNA from host cells or other microorganisms will be subject equally to amplification following annealing of the random primers and will result in a heterogenous and irreproducible amplification pattern. The requirement for small quantities of pure DNA makes this method particularly suitable for slow-growing organisms such as the mycoplasmas. The quantity of primer used is a critical parameter. It has been found that 400 ng is optimal for precise, informative pattern amplification. Less primer may result in fewer bands and more primer may result in a pattern that is too complex to accurately evaluate. The length of the primer should be short, i.e., 10-mer, and should be chosen in accordance with the G + C ratio of the organism to be studied. The typical size range for an amplified array of fragments is between 0.2 and 3.0 kb. Our studies have indicated that the use of other DNA polymerases such as the more thermostable Vent (Perkin-Elmer) and Stoffel fragment (Perkin-Elmer) generate reproducible arrays of amplified fragments, but are different from those obtained with AmpliTaq. For this reason it is recommended that the user choose a type and source of DNA polymerase and remain consistent in its use. Our laboratory has demonstrated the utility of RAPD fingerprinting for distinguishing strains of Mycoplasma gallisepticum. Ten-base oligonucleotide primers were used individually to prime the synthesis of strain-specific and highly reproducible arrays of DNA fragments from genomic DNAs of M. gallisepticum isolates (Figs. lA and IB) (Geary et aL, 1994). The results of our RAPD analysis indicate that strains of M. gallisepticum are genetically heterogenous. Mycoplasma species exhibit genotypic variability by such mechanisms as genomic rearrangements, deletions, and insertions of repetitive elements into the genome (Taylor et aL, 1988; Yogev et aL, 1991). This results in highly variable surface protein and lipoprotein antigen expression, shown to be quite common in mycoplasmas. Despite this fact, the pattern of RAPD-amplified fragments for the same strain over numerous passages is stable and reproducible. This is as would be expected, even for an organism with a high frequency of genomic rearrangements. A rearrangement breakpoint would have to fall between primer binding sites or a segment would have to be deleted in order to affect the pattern. Detecting such a change is highly unlikely considering that the average number of fragments in a typical array is approximately 10, all of which are less than 3 kb in size. Therefore, the total DNA amplified is no greater than a maximum of 20-30 kb which represents approximately 2-3% of the average mycoplasma genome. In addition to the diagnostic value of RAPD analysis, this procedure is useful in a research capacity as well (Wong and McClelland, 1994). We have applied
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Steven ). Geary and Mark H. Forsyth
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Fig. 1. DNA banding patterns of Mycoplasma gallisepticum strains following RAPD amplification with (A) primer 1281 or (B) primer 1254. The individual gel lanes were reordered via computer imaging for ease of visual comparison.
this approach to the investigation of differentially expressed genes in M. pneumoniae. Preliminary experiments were designed to determine if an upregulation of mRNA transcripts following exposure to UV irradiation could be detected. Following exposure of 1 x lO^/ml M. pneumoniae PI1428 suspended in phosphate-buffered saline to 7000 jxW/cm^ for either 0 (control) or 60 seconds, total RNA was extracted. Reverse transcriptase PCR was performed using the
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Gene Amp RNA PCR kit (Perkin-Elmer), according to manufacturer's protocols, to generate a cDNA copy of mRNA transcripts. The random primer designated SGI22 was utilized in a standard PCR amplification reaction as described earlier. Samples of each were electrophoresed in an agarose gel containing ethidium bromide. After exposure to UV, an apparent upregulation was discovered, as evidenced by the appearance of a new 1845-bp fragment band, indicating induction of transcription of the gene containing this fragment. Additional data are needed to determine the identity of this inducible gene; however, this demonstrates the applicability of this procedure as a research tool for the analysis of differentially expressed genes in mycoplasmas.
References Akopyanz, N., Bukanov, N. O., Westblom, T. U., and Berg, D. E. (1992a). PCR-based RFLP analysis of DNA sequence diversity in the gastric pathogen Helicobacter pylori. Nucleic Acids Res. 20, 6221-6225. Akopyanz, N., Bukanov, N. O., Westblom, T. U., Kresovich, S., and Berg, D. E. (1992b). DNA diversity among clinical isolates of Helicobacter pylori detected by PCR-based RAPD fingerprinting. Nucleic Acids Res. 20, 5137-5142. Geary, S. J., Forsyth, M. H., Aboul Saoud, S., Wang, G., Berg, D. E., and Berg, C. M. (1994). Mycoplasma gallisepticum strain differentiation by arbitrary primer PCR (RAPD) fingerprinting. Mol. Cell. Probes 8, 11-316. Kersulyte, D., Woods, J. P., Keath, E. J., Goldman, W. E., and Berg, D. E. (1992). Diversity among clinical isolates of Histoplasma capsulatum detected by polymerase chain reaction with arbitrary primers. J. Bacteriol. 174, 7075-7079. Micheli, M. R., Bova, R., Calissano, P., and D'Ambrosio, E. (1993). Random amplified polymorphic DNA fingerprinting using combinations of oligonucleotide primers. BioTechniques 15, 388-389. Taylor, M. A., Ferrell, R. V., Wise, K. S., and Mcintosh, M. A. (1988). Reiterated DNA sequences defining genomic diversity within the species Mycoplasma hyorhinis. Mol. Microbiol. 2, 665672. Wang, G., Whittman, T. S., Berg, C , and Berg, D. (1993). RAPD (arbitrary primer) PCR is more sensitive than multilocus enzyme electrophoresis for distinguishing related bacterial strains. Nucleic Acids Res. 21, 5930-5933. Welsh, J., and McClelland, M. (1990). Fingerprinting genomes using PCR with arbitrary primers. Nucleic Acids Res. 18, 7213-7218. Welsh, J., Pretzman, C , Postic, D., Saint Girons, I., Baranton, G., and McClelland, M. (1992). Genomic fingerprinting by arbitrarily primed polymerase chain reaction resolves Borrelia burgdorferi into the distinct phyletic groups. Im. J. Syst. Bacteriol. 42, 370-377. Williams, J. G. K., Kubelik, A. R., Livak, K. J., Rafalski, J. A., and Tingey, S. V. (1990). DNA polymorphisms amplified by arbitrary primers are useful as genetic markers. Nucleic Acids Res. 18, 6531-6535. Wong, K. K., and McClelland, M. (1994). Stress-inducible genes of Salmonella typhimurim identified by arbitrary primed PCR of RNA. Proc. Nad. Acad. Sci. USA 91, 639-643. Yogev, D., Rosengarten, R., Watson-McKown, R., and Wise, K. S. (1991). Molecular basis of mycoplasma surface antigenic variation: A novel set of divergent genes undergo spontaneous mutation of periodic coding regions and 5' regulatory sequences. EMBO J. 10, 4069-4079.
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SECTION
B
Immunological Tools
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B1 INTRODUCTORY REMARKS Joseph G. Tully
Serology still plays an important role in the diagnosis of mollicute infection and in the detection and identification of the organisms. The biologic properties of mollicutes, particularly their limited biosynthetic capabilities, a requirement for complex media, and a significandy prolonged growth phase, have greatly limited the application of many of the conventional direct cultural techniques for rapid and specific diagnoses of clinical bacterial infections. A variety of serologic tests, utilizing essentially membrane antigens of the organism, dominate the measurement of the host's immune response to mollicutes and identification of the organism to the species level. Many of the serological methods published in the earlier series of "Methods in Mycoplasmology" (Razin and Tully, 1983; Tully and Razin, 1983) are still applicable. Basic techniques for the preparation of antigens and antisera also have not changed materially, although adjuvants that rely on a metabolizable oil base (such as squalene) have been recommended to replace the earlier mineral oil-based adjuvants (Tully, 1993). Growth inhibition, metabolism inhibition, deformation, and immunofluorescence tests described earlier are utilized extensively today in the laboratory identification and speciation of mollicutes. Likewise, many laboratories continue to use the complement fixation, metabolism inhibition, immunofluorescence, and some enzyme-linked immunosorbent assays (ELISA) to measure specific antibody responses to mollicutes in humans and in other animal hosts. However, several advancements in the refinement and application of the ELISA procedure to mollicutes have been reported. Contributions in this section emphasize some of these newer techniques. As was noted in an earlier description of ELISA tests (Cassell and Brown, 1983), the technique also has a distinct 89 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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advantage in that it can be designed to measure immune responses to all three major antibody classes (IgG, IgM, and IgA) in a host. Chapter B2 covers application of the ELIS A to the immunological detection of mycoplasmal infections in a variety of small animals, laboratory rodents, and birds. With reductions in the occurrence of mycoplasmas in many commercial sources of both rodents and birds, these tests can provide a simple, rapid, and low-cost alternative in monitoring the occurrence and spread of infections, particularly those disseminated through the respiratory route. Chapter B3 describes the application of ELISA to mycoplasmal antibody detection in several important large animal infections and to an antigen capture technique, using both the ELISA and an immunoblotting procedure for the indirect detection of mycoplasmas. Although it is agreed that these serologic procedures are a significant improvement over earlier methodology, particularly for bovine and caprine pleuropneumonia, the contagious nature of these diseases and their important economic role require that much more be done to improve and standardize serologic techniques for detecting and controlling animal mycoplasma infections. One of the new advancements in the serologic analysis of mycoplasmas of human origin has been the development of an ELISA using lipid-associated membrane protein antigens (Chapter B4). These tests have demonstrated a surprisingly high degree of specificity in measuring antibody to several Mycoplasma species (M. penetrans, M. genitalium, and M. fermentans) which have been difficult to cultivate under the usual laboratory conditions. The new ELISA provides an important means to evaluate host responses to a group of mycoplasmas increasingly associated with various sexually transmitted diseases, including acquired immunodeficiency syndrome (AIDS) (see also Chapters D3 and D5, this volume). ELISA tests for respiratory tract infections are discussed extensively in Chapter B5, as well as in Chapter D2, where the diagnosis of M. pneumoniae infection is reviewed. The use of monoclonal antibodies in the ELISA has expanded the sensitivity and specificity of detection of mycoplasma antigens in biological material and in cultures. Several examples of the general approach to the use of monoclonal antibodies in ELISA testing for animal or plant infections are described in Chapter B6, as well as in several sections of Chapter B3. However, one should be aware of those special circumstances where increased specificity of a monoclonal antibody-based ELISA for antigen detection might not recognize wild-type strains of the species, such as with Spiroplasma kunkelii in plant infections (see Whitcomb, 1989) or in those Mycoplasma species that exhibit considerable phase variation in membrane antigens (see Chapter C3 in Vol. I). In these circumstances, an ELISA with a polyclonal antibody probably offers a better approach to indirect detection of the organism. The immunoblotting or immunobinding techniques, which allow individual proteins in complex mixtures to be detected and analyzed, have been of great
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importance in delineating mycoplasmal antigens (including adhesins) that function as important immunogens in a variety of hosts. Some of the current applications of these techniques to mollicutes are outlined in Chapter B8. Immunofluorescence techniques have also had wide acceptance within mollicute research. The agar plate immunofluoresence test described in the earlier methodology series (Gardella et al., 1983) still retains its dominance as the most useful serologic technique for rapid species identification of mollicutes (TuUy, 1993). In Chapter B7, a microimmunofluorescence procedure for measuring mycoplasma antibody is presented. This application has a number of advantages, especially regarding sensitivity and specificity: test readings are not affected by antibiotics in the serum, the test antigen can be standardized, and different antibody classes can be measured. Finally, an adaptation of the agar plate immunofluorescence test for laboratory distinction of M. genitalium from M. pneumoniae is outlined in Chapter B9. The need for this technique arose when M. genitalium strains mixed with M. pneumoniae were isolated from the human respiratory tract. With specific conjugated antiserum to each organism, laboratory identification and separation of mixed species of these organisms, or other mollicutes, can be accomplished.
References Cassell, G. H., and Brown, M. B. (1983). Enzyme-linked immunosorbent assay (ELISA) for detection of anti-mycoplasmal antibody. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.), Vol. 1, pp. 457-469. Academic Press, New York. Gardella, R. S., Del Giudice, R. A., and TuUy, J. G. (1983). Immunofluorescence. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.), Vol. 1, pp. 431-439. Academic Press, New York. Razin, S., and TuUy, J. G., eds. (1983). "Methods in Mycoplasmology," Vol. 1, Academic Press, New York. TuUy, J. G. (1993). Serological identification of mollicutes. In "Rapid Diagnosis of Mycoplasmas" (I. Kahane and A. Adoni, eds.), pp. 121-130. Plenum, New York. TuUy, J. G., and Razin, S., eds. (1983). "Methods in Mycoplasmology," Vol. 2. Academic Press, New York. Whitcomb, R. F. (1989). Spiroplasma kunkelii: Biology and ecology. In "The Mycoplasmas" (R. F. Whitcomb and J. G. TuUy, eds.). Vol. 5, pp. 487-544. Academic Press, San Diego, CA.
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B2 ELISA IN SMALL ANIMAL HOSTS, RODENTS, AND BIRDS M. B. Brown^ J. M. Bradbury^ and J. K. Davis
General Introduction
Basic ELISA Technology
Although enzyme-Hnked immunoassay (ELISA) has been developed for both antigen and antibody detection, this chapter is restricted to applications for detecting a specific antibody. In its simplest form, the ELISA is performed by allowing the mycoplasma antigen to absorb nonspecifically to a solid phase (usually a 96-well microtiter plate), followed by the blocking of unbound protein sites and the sequential addition (with adequate washing between reagents) of the serum sample, the secondary conjugated antibody, and the enzyme substrate. The assay is completed by developing the colored product of the enzyme reaction, and reactions are read either on a spectrophotometer or by visual inspection. Although the "kit" forms of mycoplasmal ELIS As are usually interpreted subjectively, the use of semiquantitative data obtained by determining the intensity of the color change spectrophotometrically provides better discrimination of positive and negative results. The latter application is especially important in monitoring barrier-maintained animal colonies. A semiquantitative assay is relatively inexpensive and can easily be established in any laboratory with a spectrophotometer. Recent modifications of the ELISA technique include dot-blot ELISA proce93 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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dures which rely on nitrocellulose or other membranes as the solid support. Results are usually recorded visually, although scanning densitometers can provide more quantitative data. Considerable variation in ELISA techniques exists among laboratories working with moUicutes, including differences in mollicute strains selected as antigens, antigen preparation methods, solid-phase supports, reagents, and incubation times and temperatures. All of these factors affect the test and can lead to a lack of agreement in test results among laboratories. In most cases, high quality, commercially available, conjugated secondary antibodies, substrates, and developing reagents have eliminated the need for preparation of these reagents. Availability of standard reference serum reagents for quality control purposes could also reduce variability among laboratories. This is especially important when the ELISA is used for the clinical diagnosis of mycoplasmosis. Use of ELISA in Clinical and Experimental Laboratories
ELISA methodology for the detection of specific antibody to Mycoplasma and Ureaplasma species has proven to be a useful tool in the clinical diagnostic laboratory as well as the research laboratory. In a clinical setting, antibody detection is most often used to confirm exposure to the infectious agent. This is especially true in both poultry and laboratory rodent populations where the establishment of mycoplasma-free populations is a priority. Since individual animals are rarely followed in rodents or poultry, except under experimental circumstances, monitoring of acute and convalescent titers in individual animals is rare and of limited use clinically. However, development of specific antibody can be used as a tool for seroepidemiology in monitoring the spread of disease in populations and in providing additional understanding of the dynamics involved in the interaction between the host and the infectious agent. Changes in antibody levels could precede the appearance of clinical disease and provide an early warning of potential disease outbreaks in various populations. This could be especially important in wild animal hosts, where little is known of the natural occurrence of mollicutes or their possible role in disease. In experimental situations, ELISA technology is particularly useful. The ELISA is flexible in that both serum and secretions can be analyzed for specific antibody. Not only can seroconversion be monitored, but immunoglobulin classspecific responses can be determined. Because of differences in biological activity, the presence or the induction of specific subclasses may be of particular interest. ELISA in Laboratory Rodent Infections
Traditional methods of serologic diagnosis of mycoplasmal infections, such as complement fixation, hemagglutination, hemagglutination inhibition, and meta-
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bolic inhibition tests, have been of little use in laboratory rodents because they are relatively insensitive and depend on antibody function. In well-managed, barrier-maintained rodent colonies, both the prevalence of infected animals and the numbers of organisms per animal are usually low. This situation makes detection by serologic methods very difficult. ELISA for mycoplasmal infections in rodents has proven to be the most suitable serologic test for screening large numbers of rodents (Cassell et al, 1986; Davidson et al., 1981, 1994). A comprehensive review of all aspects of the diagnosis of rodent mycoplasmosis has appeared (Davidson et al., 1994). The test is relatively inexpensive to run, is sensitive, and gives rapid results. In addition, animals do not have to be killed to obtain enough blood for the test. Blood may be collected from anesthetized animals by tail or orbital bleeding. However, the assay currently available cannot distinguish among Mycoplasma species, and quality control of the assay, as with all serological assays, is critical for reliable results. ELISA in Poultry and Other Avian Infections The most common method of diagnosis of either M. gallisepticum or M. synoviae infection in poultry is determination of antibody status. The serum plate agglutination test is commonly used to screen for mycoplasma infections but has been associated with false positives. In the United States, both hemagglutination inhibition and ELISA are approved as confirmatory tests by the National Poultry Improvement Program, but their use also must be specifically approved by each state. Several commercial ELISA kits are available on the market for detecting antibodies to M. gallisepticum and M. synoviae in chicken sera. At least one company markets such kits specifically for use with turkey sera, as well as a kit for detecting M. meleagridis antibodies in turkeys (KPL Inc., Gaithersburg, MD). One commercial kit is designed to react with antibodies to either M. gallisepticum ox M. synoviae (IDEXX Laboratories Inc., Westbrook, ME). Recent advancements in the preparation of ELISA antigens, with removal of nonspecific antigenic components, have improved specificity and decreased cross-reactions. ELISA tests using affinity-purified antibodies are available with good results for both M. gallisepticum and M. synoviae. However, there is no suitable ELISA for detecting M. iowae, an organism found mainly in turkeys. This mycoplasma does not appear to elicit a consistent humoral antibody response, and experimental studies have shown an unacceptably high number of falsepositive reactions in normal turkey sera (Jordan et al., 1987). A "blocking ELISA" has been proposed for detecting M. gallisepticum infection (Czifra et al., 1993). The test uses a monoclonal antibody (MAb) which recognizes an epitope on a 56-kDa polypeptide that appears to be permanently expressed on the mycoplasma. In this system, ELISA plates are coated with M. gallisepticum whole cell antigen, and the serum under test is added undiluted
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(Diagnosticum, Budapest, Hungary; Svanova Biotech, Uppsala, Sweden). The reaction is assessed by the extent of blocking that occurs when the conjugated MAb is added. One assay can be used for sera from any host species without adaptation as long as the relevant antigen is recognized by the individual animal. ELISAs have also been used successfully to detect M. gallisepticum antibodies in diluted or extracted egg yolk (Mohammed et al., 1986; Brown et al., 1991). When the currently available commercial M. gallisepticum ELISAs were compared with other commonly used serological tests, i.e., the rapid serum agglutination (RSA) and hemagglutination inhibition (HI) tests, the RSA detected antibodies slightly earlier than the ELISA or HI tests (Kempf et al., 1994; J. M. Bradbury and J. Lewis, unpublished observations). However, some poultry companies find ELISAs more convenient for the routine screening of large numbers of birds because they already employ ELISA technology in screening for the many important poultry viruses. As with other mycoplasma ELISAs, the sensitivity of the avian mycoplasma ELISAs is determined to some extent by the recommended cutoff levels for positive and suspicious reactions. The sensitivity in some cases may be deliberately "dampened down" to avoid the well-recognized cross-reaction between M. gallisepticum and M. synoviae. This is a dangerous practice as it inherently implies the assay conditions have not been optimized and low level infections may be missed. The development of ELISA antibodies may be suppressed if the birds have received antibiotic treatment. In addition, a serological response will not differentiate birds that have been vaccinated against mycoplasmal infection from those naturally infected. Also, it should be noted that polymerase chain reaction (PCR)-based tests have been developed and these should be evaluated as well as serological tests for their efficacy in detecting infection (see Chapter D9, this volume). EUSA in Other Small Animals
For the most part, serological assays have been limited to poultry and rodents, although mycoplasmal infections in populations of environmentally threatened wild tortoise populations have been examined using ELISA (Schumacher et al., 1993). In addition, antibody to both M. gallisepticum and M. synoviae has been detected in sera from golden conures (psittacine) located in a private zoological collection (M. B. Brown, unpublished data). The major deterrent to the development of ELISAs for other animals is the availability of a specific antibody which recognizes the animal species. The competitive (blocking) ELISA could be used with any species, but it is imperative that the antigen(s) or epitopes recognized by the control serum are also recognized by all infected animals. Commercial conjugates generally are available for most domestic species. Conjugates for exotic animals may be prepared commercially by specialized companies or by university facilities, such as the Biotechnologies for the Ecological,
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Evolutionary, and Conservation Sciences Immunological Analysis Laboratory at the University of Florida. Alternatively, a blocking ELISA can be developed.
Antigen Preparation Antigen preparation is a critical step for the success of any ELISA. Antigen preparations may consist of whole cells, lysates, partially purified protein preparations, purified proteins, or even recombinant proteins. Regardless of how the antigen is prepared, certain requirements should be applied. First, the proteins present in the antigen preparation should be present on all (or at least most) field strains of the mycoplasmal species. The preparative steps should be standardized to assure batch to batch reproducibility of the antigen. Given the documented antigenic variation of many mollicutes, it is imperative that purified proteins or recombinant proteins used as antigens should be constitutively expressed and recognized by infected animals at all stages of the infection. If purified proteins are used as antigens, it is important to use a mixture of proteins rather than reliance on a single protein. One of the most common antigen preparations used is a lysate of whole mycoplasma cells to produce the antigen for the assay (Horowitz and Cassell, 1978). This method of antigen preparation is used because it is easily accomplished and yields a reliable, stable antigen for routine use. This general method may be used for most mollicutes, but the harvesting times and protein yield vary considerably from mollicute species to species. Antigens prepared by this method are stable at -70°C for at least 6 months. The effect of storage conditions on antigenicity must be assessed for each antigen preparation and for each mollicute species. Several other ELISA antigen preparation methods have been reported and are described elsewhere in this volume (see Chapter B3). The following technique will usually provide a satisfactory antigen. 1. Grow the test organism in 1-3 liters of broth medium until the culture reaches the peak logarithmic phase. Harvest the organisms by centrifugation (10,000 to 20,000 g), 2. Wash the organisms three times in sterile phosphate-buffered saline (PBS, pH 7.4) by adding PBS at a ratio of 10 times the volume of the pellet, mixing well, and then centrifuging the suspension. Pour off the supernatant. Repeat the wash once again. Resuspend the pellet in a small volume (1 to 10 ml) of PBS. 3. Transfer a small amount of the final suspension into beef heart infusion broth (or other suitable medium) to check for bacterial contamination. Note that some mycoplasmal species will sometimes grow on 5% sheep blood agar plates. 4. Determine the protein concentration of the final suspension. 5. Dilute the suspension in sterile PBS to a final concentration of 5 mg protein/ml. Store aliquots of the suspension at -70°C.
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6. Dilute a small amount of the 5-mg/ml suspension to 1:100 (0.05 ml of suspension to 0.95 ml PBS) and read absorbance at 540 nm. 7. Dilute the remainder of the suspension 1:20 in carbonate-bicarbonate buffer (0.05 M, pH 10.0). Incubate the diluted suspension at 37°C. 8. Check the absorbance of the diluted suspension at 5-minute intervals until the absorbance of the suspension is 50% of the absorbance of the PBS suspension in step 6. This usually takes about 15 minutes. 9. Stop the lysis by adding 2.2 g of boric acid per 100 ml of suspension. 10. Determine the protein concentration of the lysate, and store aliquots at -TOT.
Conjugate and Substrate The availability of high quality conjugated antisera has increased dramatically since the 1980s. Comprehensive listings of anti-immunoglobulins are available in publications such as "Linscott's Directory." If the host species is not commercially available, many private companies or university core facilities can prepare custom antisera at a nominal charge. The most common enzymes used for conjugation to anti-immunoglobulins are alkaline phosphatase and horseradish peroxidase; biotinylation also is common. Although less common in ELISA, additional enzymes including urease and 3-galactosidase or fluorochromes can be used. Most antiimmunoglobulins, especially IgG, are available already labeled, and subclassspecific antibodies are readily available for rodent immunoglobulins. If not available, several methods and commercial labeling kits are available (Harlow and Lane, 1988). In addition to specific anti-immunoglobulins, some assays use labeled protein G or protein A to detect bound immunoglobulins. It should be noted that these proteins may vary in their ability to bind IgG from different host species and the binding capacity for the host IgG of interest should be established if this choice is made. A variety of substrates are available for the enzymes just discussed. For the purposes of this discussion, only soluble substrates will be considered. Insoluble substrates are available for dot-blot or other detection systems which immobilize antigens on nitrocellulose or polyvinyl membranes. Most substrates and their buffers are available from commercial sources. For alkaline phosphatase, the substrate of choice is p-nitrophenyl phosphate (PNPP) which yields a yellow solution. The reaction is stopped by the addition of 3 N NaOH, and the test is read at 405 nm. The substrate buffer is described elsewhere in this volume (see Chapter B5). For horseradish peroxidase, the substrates of choice are 2,2'azinobis (3-ethylbenzothiazoline-6-sulfonic acid (ABTS), o-phenylenediamine
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dihydrochloride (OPD), or 5-aminosalicylic acid (5AS). ABTS yields a bluegreen color, with the reaction stopped by the addition of 1% sodium dodecyl sulfate and the test read at 405 nm. OPD yields an orange-brown product and is read at 492 nm; sensitivity increases two- to fourfold if the reaction is stopped with 3 N HCl or 3 M H2SO4 and the test is read at 450 nm. 5AS yields a brown color which is read at 450 or 550 nm, after the reaction is stopped with 3 A^ NaOH.
Assay Standardization In developing an ELISA there are numerous factors that can affect results and their interpretation. The first decision is whether or not a single dilution, or multiple dilutions, of the specimen should be tested. Results may be reported as absorbance, end point titration, significant rises in titer, activity as compared to some "standard" serum, or simply positive, suspect, or negative when compared to a known positive or negative serum sample. A single serum sample often is adequate for diagnostic purposes. Because single dilution ELISA assays are heavily dependent on antibody affinity, quantitative standardization in terms of milligrams of antibody present is almost impossible. Although serum is the most common specimen examined, egg yolks have been effectively used in avian and reptile systems (Mohammed et aL, 1986; Brown et al., 1991; M. B. Brown and I. M. Schumacher, unpublished data). As a screening tool in the poultry industry, egg yolk testing may offer an advantage under some circumstances because the eggs can be readily collected, thus avoiding the time-consuming and invasive procedures of drawing blood. Lavages from respiratory and genital tracts have been used, but these are used most often in experimental situations where serum antibody levels are also examined (Simecka and Cassell, 1987; Brown and Reyes, 1991; Elfaki et aL, 1992). In selecting the dilutions of antibody and reagents, it is important that the amount of antiserum in the unknown test sample should be the only limiting factor. Adjusting the concentration of antigen, serum dilution, and conjugate dilution by simultaneous checkerboard titration is likely to cause trouble and often leads to a choice of conditions where some factor besides the specific antibody in the test serum is limiting. Choosing Plates The first step in establishing the assay is choosing an acceptable solid-phase support, which is usually a microtiter plate. The mean, standard deviation.
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coefficient of variation, and ratio of immune to normal serum absorbance of the colored product of the enzyme reaction should be determined for each sample. The coefficient of variation should be as small as possible, and the ratio of the immune to normal serum should be as large as possible. Most plates show a significant "edge effect," and avoidance of the outside rows of wells for the test is advisable. The criteria used for selecting a microtiter plate are a coefficient of variation less than 0.5 to 0.75 for both immune and normal serum and the highest possible ratio between hyperimmune and normal serum. Blocking Solutions In almost all ELISA assays, unbound sites that can nonspecifically absorb either immunoglobulin in the sample or conjugated secondary antibody remain on the microtiter plates. These unbound sites should be blocked by inert protein. A variety of blocking solutions have been used, including Tween 20, BLOTTO, 2% w/v bovine serum albumin (BSA), 5% w/v BSA, 2% v/v fetal calf serum (PCS), and 5% v/v PCS. The ability to block both the binding of normal serum and the binding of the conjugated secondary antibody to the microtiter wells should be assessed. The choice of the blocking reagent to be used must be examined for each microtiter plate and for each antigen preparation. Determining Antigen Concentration Dilutions of antigen, test serum, and conjugate can be determined as follows. Arbitrarily select a dilution of a known immune serum and the conjugated secondary antibody that will give a high concentration of these two reagents so that they cannot be the limiting reagents in the assay. Coat the microtiter plates with various concentrations (0.1 to 50 |JLg/ml of protein) of mycoplasma antigen. Run the assay in the routine manner. Plot the absorbance versus the antigen concentration. Pick an antigen concentration that is well on the plateau of the absorbance curve for all future tests. A protein concentration of 2-10 |xg/ml is usually adequate to ensure that the antigen is never the limiting factor in the assay. Determining Conjugate Concentration The working concentration of each conjugate is determined by reacting antigen-coated wells with PBS and then using dilutions (1:500 to 1:10,000) of the affinity-purified conjugates in the assay. Plot the absorbance versus the dilution of the conjugate and choose the lowest dilution that does not show appreciable nonspecific binding to the antigen. As a general rule, the conjugate dilution should be less than 1:5000 to ensure that the conjugate is never the limiting factor in the assay.
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Determining Serum Dilution
The serum dilution to use is the last of these parameters to be determined because the working concentrations of antigen and conjugate and which microtiter plate and blocking reagent works best should already be established. If multiple dilutions and end point titration are used, then this is not necessary. The plates are coated with antigen, blocked, and reacted with different concentrations of serum (dilutions range from 1:10 to 1:1500; lavages or egg yolk may be used as well), followed by reaction with the conjugate. Plot the absorbance vs the serum dilution. Choose the lowest dilution that shows little nonspecific absorption to the plates. Sometimes a compromise must be reached at this step because a very low nonspecific absorbance and a low serum dilution are both desirable. The former is necessary for maximum sensitivity, but the latter is required to allow the detection of small amounts of antibody. As a final check of the reaction reagents, the conjugate titration should be run again with normal and immune sera as well as the PBS control. When the absorbance is plotted versus the conjugate dilution, the normal serum and PBS control curves should be virtually identical. There should also be a wide separation between the values of the immune serum and the normal serum at the conjugate dilution selected for use in the assay. Incubation Times
A wide variety of incubation conditions have been reported for ELISAs. In some cases, incubation times and temperatures are adjusted for convenience (i.e., overnight incubations or room temperature vs 37°C). Ideally, the conditions for each incubation step should be determined by similar methods to those used to determine which microtiter plates were acceptable for use in the assay. The mean, standard deviation, coefficient of variation, and ratio of the absorbance of the immune to normal serum are determined for each incubation procedure. The coefficient of variation should be as small as possible, indicating reproducibility. Likewise, the ratio of immune to normal serum absorbance values should be as large as possible, indicating the range of the assay and its potential sensitivity. In addition, the ratio of the normal serum to the PBS control values should be determined because this is a measure of the nonspecific activity in the system. This ratio should be as close to 1 as possible, thereby indicating no nonspecific activity.
Quality Control One of the disadvantages of the ELISA assay is its extreme sensitivity to changes in procedure. If a minor variation is inadvertently introduced into the
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procedure at any one point, this variation may be amplified in the test results. Therefore, stringent quality control measures are required. The Youden plot has been proven to be very useful (Jeffcoate, 1982). This method requires the use of dilutions of immune serum simulating high, mid-range, and low concentrations of specific antibody to the antigen in question. To determine within run and between run variability in the assay, one sample of each dilution is tested at the beginning of a set of tests and another is included at the end of the assay. The absorbance values obtained on these control samples are plotted against each other, resulting in a linear plot with a slope of 1 at a 45° angle from the X and Y axes. Variability within an assay is indicated by the spread of points away from the 45° line. The maximum divergence from the line should be no more than a 10% deviation. Day to day variation is indicated by the spread of values away from the averages of the last 10 assay runs. If a value for an individual day deviates by more than 10% from the average of the values for the last 10 runs, the run should be repeated. Other indications that a run needs to be repeated are when all three samples fall on one side of the line and are outside of a 5% deviation range.
Limitations and Other Considerations ELISA has many advantages, but it depends on the ability of the animal to produce a specific antibody. It is important to understand the natural history of the host immune response to assess the efficacy of ELISA. For example, in rodents there is a delay in antibody production during subclinical infections, resulting in a 1- to 3-month "lag time" when infected animals cannot be detected by serologic methods (Cassell et aL, 1986). However, on farms with chickens of various ages, the number of seropositive birds increases exponentially with the age of the bird, suggesting a rapid seroconversion to avian mycoplasmas (Brown etal. 1991). In rodents, a crucial problem is cross-reactions among the murine mycoplasmas (Minion et aL, 1984; Cassell et aL, 1986; Davis et aL, 1986; Watson et aLy 1987). All of the murine mycoplasmas share some common antigens; these appear to be particularly troublesome in trying to discriminate M. pulmonis infection from M. arthritidis or M. muris infection (Cassell et aL, 1986; Watson et aL, 1987; Davidson et aL, 1994). In poultry, the inability to differentiate between infected birds and vaccinated birds may also be troublesome. As ELISAs are developed for mycoplasmal infections in other hosts (i.e., cats, dogs, tortoises), the potential for cross-reactive antigens should be of some concern. Such antigens may also pose a particular problem in ELISA tests on exotic animals and other hosts with less well-defmed mycoplasmal flora.
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Optimum serologic detection of mycoplasmal infections depends on the sensitivity and specificity of the test system and on the interpretation of the results obtained. Technological advancements now suggest the possibility that serologic tests may eventually be replaced by a variety of antigen detection methods, including DNA hybridization methods with specific gene probes or the polymerase chain reaction (Kleven et aL, 1988; Lauerman et aL, 1993). Ultimately, stringent field testing combined with careful experimental infection studies will be required to define the reliability of such diagnostic procedures.
References Brown, M. B., and Reyes, L. (1991). Immunoglobulin class- and subclass-specific responses to Mycoplasma pulmonis in serum and secretions of naturally infected Sprague-Dawley female rats. Infect. Immun. 59, 2181-2185. Brown, M. B., Stoll, M. L., Scasserra, A. E., and Butcher, G. D. (1991). Detection of antibodies to Mycoplasma gallisepticum in egg yolk versus serum samples. J. Clin. Microbiol. 29, 29012903. Cassell, G. H., Davis, J. K., Cox, N. R., Brown, M. B., and Minion, F. C. (1986). State of the art detection methods for rodent mycoplasmas. In "Complications of Viral and Mycoplasmal Infections in Rodents to Toxicology Research and Testing" (T. E. Hamm, ed.), pp. 143-160. Hemisphere Press, Washington, DC. Czifra, G., Sundquist, B., Tuboly, T., and Stipkovits, L. (1993). Evaluation of a monoclonal blocking enzyme-linked immunosorbent assay for the detection of Mycoplasma gallisepticumspecific antibodies. Avian Dis. 37, 680-688. Davidson, M. K., Lindsey, J. R., Brown, M. B., Schoeb, T. R., and Cassell, G. H. (1981). Comparison of methods for detection of Mycoplasma pulmonis in experimentally and naturally infected rats. J. Clin. Microbiol. 14, 646-655. Davidson, M. K., Davis, J. K., Gambill, G. P., Cassell, G. H., and Lindsey, J. R. (1994). Mycoplasmas of laboratory rodents. In "Mycoplasmosis in Animals: Laboratory Diagnosis" (H. W. Whitford, R. F. Rosenbusch, and L. H. Lauerman, eds.), pp. 97-133. Iowa State Univ. Press, Ames. Davis, J. K., Cassell, G. H., Gambill, G., Cox, N., Watson, H., and Davidson, M. (1986). Diagnosis of murine mycoplasmal infections by enzyme linked immunosorbent assay (ELISA). Isr. J. Med. Sci. 23, 717-722. Elfaki, M. G., Ware, G. O., Kleven, S. H., and Ragland, W. L. (1992). An enzyme-linked immunosorbent assay for the detection of specific IgG antibody to Mycoplasma gallisepticum in sera and tracheobronchial washes. J. Immunoassay 13, 97-126. Harlow, E., and Lane, D. (1988). "Antibodies: A Laboratory Manual," pp. 319-358. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Horowitz, S. A., and Cassell, G. H. (1978). Detection of antibodies to Mycoplasma pulmonis by an enzyme-linked immunosorbent assay. Infect. Immun. 22, 161-170. Jeffcoate, S. L. (1982). Use of Youden plot for internal quality control on the immunoassay laboratory. Ann. Clin. Biochem. 19, 435-437. Jordan, F. T. W., Yavari, D., and Knight, D. L. (1987). Some observations on the indirect ELISA for antibodies to Mycoplasma iowae serovar I in sera from turkeys considered to be free from mycoplasma infections. Avian Pathol. 16, 307-318.
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Kempf, I., Gesbert, F., Guittet, M., Bennejean, G., and Stipkovits, L. (1994). Evaluation of two commercial enzyme-linked immunosorbent assay kits for the detection of Mycoplasma gallisepticum antibodies. Avian Pathol. 23, 329-338. Kleven, S. H., Morrow, C. J., and Whithear, K. G. (1988). Comparison of Mycoplasma gallisepticum strains by hemagglutination-inhibition and restriction endonuclease analysis. Avian Dis. 32, 731-741. Lauerman, L. H., Hoerr, F. J., Sharpton, A. R., Shah, S. M., and van Santen, V. L. (1993). Development and application of a polymerase chain reaction assay for Mycoplasma synoviae. Avian Dis. 37, 829-834. Minion, F. C., Brown, M. B., and Cassell, G. H. (1984). Identification of cross-reactive antigens between Mycoplasma pulmonis and Mycoplasma arthritidis. Infect. Immun. 43, 115-121. Mohammed, H. O., Yamamoto, R., Carpenter, T. E., and Ortmayer, H. B. (1986). Comparison of egg yolk and serum for the detection of Mycoplasma gallisepticum and M. synoviae antibodies by enzyme-linked immunosorbent assay. Avian Dis. 30, 398-408. Schumacher, I. M., Brown, M. B., Jacobson, E. R., Collins, B. R., and Klein, P. A. (1993). Detection of antibodies to a pathogenic mycoplasma in desert tortoises (Gopherus agassizii) with upper respiratory tract disease (URTD). J. Clin. Microbiol. 31, 1454-1460. Simecka, J. W., and Cassell, G. H. (1987). Serum antibody and cellular responses in LEW and F344 rats after immunization with Mycoplasma pulmonis antigens. Infect. Immun. 55, 731-735. Watson, H. L., Cox, N. R., Davidson, M. K., Blalock, D. K., Davis, J. K., Dybvig, K., Horowitz, S. A., and Cassell, G. H. (1987). Mycoplasma pulmonis proteins common to the murine mycoplasmas. Isr. J. Med. Sci. 23, 442-447.
B3 ELISA IN LARGE ANIMALS J. Nicolet and J. L. Martel
Introduction Enzyme-linked immunosorbent assays (ELISA) are widely used as a diagnostic tool in mycoplasmal infections of large animals. The need for such techniques in the diagnosis of mycoplasmosis is commonly recognized, considering the economic importance of respective diseases in cattle [contagious bovine pleuropneumonia (CBPP) and mastitis], in small ruminants [contagious caprine pleuropneumonia (CCPP) and contagious agalactia], and in swine (enzootic pneumonia). For this purpose, the principal objectives are to devise efficient diagnostic tests for routine performance in field conditions available and to develop tests for the identification of mycoplasmas in culture. There are many variations in the performance of ELISAs. In large animals, the indirect ELISA for the detection of antibodies in serum and milk is most commonly used. Several new applications using monoclonal antibodies in sandwich or competitive ELISA have been introduced for detecting not only antibodies but also antigens in biological material and in culture. The goal of this chapter is to make a critical survey of the methodology of the ELISA techniques described for large animals. Antigen Preparation The quality of antigen preparations chiefly used for indirect ELISAs (capture of antibodies) is decisive for the specificity and the sensitivity of the test. A variety of different antigen preparations have been proposed. Representative 105 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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antigen preparations used in indirect or competitive ELISAs in large animals are described here. WHOLE CELLS
All antigen preparations basically start with the production of whole cells. 1. Harvest the culture of mycoplasmas grown in appropriate media, preferably at the logarithmic phase of growth, by centrifugation at 10,000 to 20,000 g for 20 to 30 minutes at 4°C. 2. Wash organisms three times in sterile phosphate-buffered saline (PBS), pH 7.4, in Tris-buffered saline (TBS), or in sterile physiological saline. 3. Resuspend the pellet in 1/10 to 1/100 of the original volume in buffer or saline and determine the protein concentration. This stock suspension can be used for additional antigen preparations. 4. If a whole cell preparation is used as antigen, adjust the protein content to 1 mg/ml, aliquot, and store at -70° or -20°C. SONICATED CELLS
1. Disrupt the suspended whole cells (stock or diluted suspension) by sonication on ice in alternating cycles of 30 seconds until the suspension is clarified. 2. Remove insoluble material by centrifugation at 10,000 g for 15 minutes at 4°C. 3. Aliquot and store at -70° or -20°C. SOLUBIUZED CELLS
Many solubilization methods using different detergents have been proposed. They are commonly prepared as a first step of purification, but they can also be used as a test antigen, e.g., after a Tween 20 solubilization (Nicolet et al., 1980). Tween 20 Solubilization Method
1. Centrifuge whole cell suspensions of known protein concentration. 2. Resuspend pellet in 0.0125 M phosphate buffer, pH 7.2, containing 1% (v/v) Tween 20 (1 ml buffer/mg protein). As an alternative, 0.025 M Tris-HCl buffer, containing 0.25 M NaCl (pH 7.5) and 2% (v/v) Tween 20, can be used. 3. Incubate for 30 minutes at 37°C in a rotary shaker water bath. 4. Centrifuge at 30,000 to 40,000 g for 30 minutes at 4°C. 5. Filter supernatant fluid (0.2-|jLm pore diameter membrane). 6. Store at -20°C or, for current use, at 4°C for up to 2 weeks.
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PURIFIED ANTIGENS
The purification of reactive antigens can be achieved by chromatographic separation of solubihzed protein antigens. The most promising results have been obtained by the procedure starting with Tween 20 solubilized antigen and with separation on a G-25 column (Nicolet et al., 1980) or Sephacryl column chromatography (Kazama et al., 1989). Sephadex G-25 Column
Chromatography
1. Place the Tween 20 antigen on a Sephadex G-25 fine (Pharmacia) colunm equilibrated with 0.0125 M phosphate buffer, pH 7.1. 2. Collect elution fractions, monitor at 280 nm, and determine the protein content and reactivity for each peak. 3. Store at — 20°C or, for current use, at 4°C for up to 2 weeks. Sephacryl S-300 Column
Chromatography
The same procedure is used as for Sephadex G-25 chromatography, but a Sephacryl S-300 (Pharmacia) column equilibrated with 0.1 M Tris-HCl buffer containing 0.5 M NaCl (pH 7.5) is used. General Methodological Considerations on Performance of Assays
Test procedures for ELISAs are established, and recommended protocols are given in different manuals. A few key points and recommendations have to be taken into consideration so as to ensure specificity and reproducibility of the tests and to allow a proper evaluation of the results (Carpenter, 1992; Wright et al., 1993). 1. The optimum concentration of antigen and conjugate dilution in noncompetitive assays should be established by checkerboard titration with highly positive and negative controls along with buffer alone. 2. For determination of antibodies in routine tests, the commonly used method is a single dilution of the test sample, varying from 1/10 to 1/200, in a blocking buffer according to the assay. In the indirect ELISA, serum concentrations of 1/100 or 1/200 are commonly tested. Milk samples may be tested in undiluted or diluted (1/2) form. 3. An internal quality control should be performed by means of selected strongly positive, weakly positive, and negative reference standards (if possible international standards).
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4. Positive and negative controls, which are representative of the local animal population, generally serve as working standards for routine diagnostic use. These controls have previously been calibrated against reference standards. 5. Reading of color after enzymatic reaction can be done visually for qualitative appreciation. For more precise and quantitative readings, a spectrophotometer is preferred. According to the substrate used, absorbance (OD) is determined at 492, 450, or 405 nm for horseradish peroxidase (HRP) and at 405 nm for alkaline phosphatase. 6. Data expression of OD mean values of duplicate samples should preferably be given in percentage of positivity relative to a strongly positive reference standard (=100%) for indirect ELISA and in percentage of inhibition of the standard competing antibody (=0%) for competitive ELISA. 7. To obtain the "cutoff" value determining negative or positive reactors, samples from representative groups of noninfected and infected animals should be tested. As the cutoff will determine the diagnostic sensitivity and specificity as well as the predictive value of the test, the selection of the respective value for a given test will depend on the application required. By using preselected sensitivity/specificity, adjustments can be made in accordance with the purpose of the test. In the ideal event, the cutoff value of a test is chosen to maximize the benefit that accrues from testing a population. To this effect, the benefit of the testing and the economic and social consequences of misdiagnoses as well as the prevalence of the disease must be taken into consideration. 8. Validation of a new assay should be performed by comparing the diagnostic performance on the same test samples with the assays in current use.
Types of Immunoassays in Diagnosis of Mycoplasmal Infections in Large Animals
A selection of different techniques in use for the diagnosis of important veterinary mycoplasmosis in large animals serves as examples for basic assay procedures. SWINE ENZOOTIC PNEUMONIA: Mycoplasma
hyopneumoniae
Several indirect ELISAs have been developed to detect antibodies in serum or in colostral milk. Most of them use a Tween 20 antigen purified by either Sephadex G-25 or Sephacryl S-300 column chromatography (see earlier discussion). Indirect ELISA for Antibody
Capture
1. Add 100 |xl of purified antigen diluted in carbonate coating buffer (pH 9.6) at an optimal concentration to a polystyrene microtiter plate.
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2. Incubate for 2 hours at room temperature or overnight at 4°C. 3. Remove antigen excess. Wash three to six times with PBS containing 0.01% Tween (PBS-T) or PBS-T with 1-2% skim milk (Difco) as a blocking agent (PBS-TS). 4. Add 100 |xl of test and control samples diluted in PBS-T or PBS-TS at a concentration of 1/100 for serum and 1/2 for milk. 5. Incubate for 90 minutes at room temperature and wash. 6. Add to each well 100 jxl of the appropriate dilution (PBS-T) of the conjugate (antiporcine IgG H+L) labeled with HRP. 7. Incubate for 90 minutes at room temperature and wash. 8. Add 100 jxl of chromogenic substrate (2 mM) diammonium-2,2'-azinobis[3-ethyl-2,3-dihydrobenzthiazole 6-sulfonate], 2.6 mM H2O2) in phosphateacetate buffer, pH 4.2. 9. Incubate at room temperature for 15 to 30 minutes. 10. Measure the absorbance at 405 nm. CONTAGIOUS AGALACTIA OF SMALL RUMINANTS: Mycoplasma agalactiae
For the contagious agalactia of goats and sheep, few reports on the development of assays for detecting antibodies are available. Satisfactory results have been recorded from tests using sonicated or Tween 20-solubilized cells as the antigen. The indirect ELISA for serum and milk is performed according to the procedure described earlier for swine enzootic pneumonia. CONTAGIOUS BOVINE PLEUROPNEUMONIA: Mycoplasma mycoides subsp. mycoides SC (SMALL COLONY FORM, Mmm SC)
The specificity of immunoassays for CBPP is drastically impaired by complex cross-reactions with other mycoplasmas belonging to the "mycoides cluster" [i.e., M. mycoides subsp. mycoides LC (large colony), M. mycoides subsp. capri, M. capricolum subsp. capricolum, M. capricolum subsp. capripneumoniae, 3.nd Mycoplasma sp. serogroup 7]. The use of monoclonal antibodies in sandwich or competitive ELISAs offers a possibility to circumvent this problem. Sandwich ELISA for Antigen
Detection
This assay detects Mmm SC in biological samples and identifies unequivocally Mmm SC in culture even after only a few days (Brocchi et al., 1993). 1. Coat microplates, overnight at 4°C, with 50 jxl of a selected pair of trapping specific monoclonal antibodies (MAbs) (10 jxl/ml) in sodium-bicarbonate buffer, pH 9.6. 2. Wash three times with PBS (pH 7.4) containing 0.05% Tween 20. 3. Add 50 |JL1 of sequential dilutions of the test antigen [liquid medium culture
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or infected tissue extract in PBS (pH 7.4) containing 0.05% Tween 20 and 1% yeast extract]. 4. Incubate for 1 hour at 37°C. Wash three times. 5. Add 25 |xl of a selected pair of pretitrated MAbs labeled with HRP. 6. Incubate for 1 hour at 37°C. Wash three times. 7. Add 50 |xl of substrate o-phenylenediamine 1% (w/v) and 0.02% H2O2 (v/v) in phosphate-citrate buffer (pH 5.0). 8. Incubate for 10 minutes at room temperature, and stop the reaction with 50 |xl of 2 M sulfuric acid. 9. Read the absorbance value at 492 nm.
Competitive
BUSA for Antibody Assessment
This assay detects specific antibodies against Mmm SC-specific epitopes avoiding cross-reactions with other mycoplasmas (Brocchi et al., 1993). The basic procedure is the same as described earlier. 1. Coat a selected pair of MAbs. 2. Add 50 |xl of an inactivated and pretitrated liquid culture (antigen) Mmm SC (strain PGl) and incubate for 1 hour at 37°C. Wash. 3. Add 50 (xl of threefold dilutions of test serum and incubate for 1 hour at 37°C. 4. Directly add 25 |xl of a competitive labeled pair of MAbs and incubate for 1 hour at 37°C. 5. Wash, add substrate, stop the reaction, and read at 492 nm. The serum titer is calculated as the dilution which competes to a 50% level for the antigen binding. CONTAGIOUS CAPRINE PLEUROPNEUMONIA: Mycoplasma capricolum capripneumoniae, syn. BIOTYPE F38
subsp.
As a member of the "mycoides cluster," M. capricolum subsp. capripneumoniae shows cross-reactivity with other members of the group. Therefore, the specificity of serological tests is questionable. Indirect ELISA
As described earlier, assays with whole cell antigens (sonicated or extracted) can basically be used as a screening test. However, other mycoplasmas, mainly M. mycoides subsp. mycoides LC and M. mycoides subsp. capri, might erroneously be taken for etiologic agents of the CCPP syndrome. In this case, the serologic results should be confirmed by the isolation of the infectious agent.
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Assay for Antigen Capture
This assay, detecting mycoplasmal antigens in pleural fluid with a detection threshold of about 10"^ organisms/ml, is proposed as a simple and rapid diagnostic tool for field diagnosis of CCPP (Guerin et al., 1993). In using a polyclonal antiserum for capture, cross-reactivity is obvious, particularly with M. mycoides LC, and confirmation by culture is required. 1. Cut small pieces of nitrocellulose membrane (MSI Nitroplus 0.22 |xm) and place them in a microtiter plate with flat-bottom wells. Avoid spoilage with finger prints. 2. Add 5 |xl of test sample (pleural fluid) to the membrane piece and allow to dry. 3. Wash for 3 minutes with 200 JJLI of Tris-buffered saline (0.05 M Tris, 0.2 M NaCl, pH 7.4) on a microplate agitator. 4. Incubate for 30 minutes at room temperature in 200 |JL1 of blocking buffer (Tris-buffered saline with 0.02% Tween 20 and 10% horse serum) with slow agitation. 5. Remove the well content and add 100 |JL1 of polyclonal rabbit immune serum in an optimal dilution (blocking buffer). 6. Incubate for 30 minutes at room temperature with slow agitation. 7. Remove fluid and wash three times for 3 minutes each in 100 |JL1 of Trisbuffered saline. 8. Dispense 100 \x\ of HRP-labeled antirabbit Ig in optimal dilution (blocking buffer). 9. Incubate for 30 minutes and wash three times. 10. Add 100 ml of substrate (0.05% 4-chloro-l-naphthol; 0.01% H2O2 in distilled water). A purple color, appearing after a few seconds, indicates a positive reaction. BOVINE MYCOPLASMAL MASTITIS: Mycoplasma
bovis
Various ELISAs have been proposed for the detection of antibodies in the diagnosis and control of M. bovis infections in cattle. Different conventional indirect ELISAs are suitable for this purpose. Recently proposed antigen capture ELISAs use monoclonal antibodies for detecting M. bovis in milk (see Chapter B6, this volume). These methods are essentially based on the procedures described for sandwich ELISA (described earlier). The limit of the detection level is in the range of 10^ CFU/ml milk (Heller et al., 1993). IDENTIFICATION OF RUMINANT MYCOPLASMAS
A dot immunobinding assay for the identification of ruminant mycoplasmas from broth culture was described (Poumarat etal, 1991). The basic procedure is
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the same as that described eariier for CCPP. The originality of the method is based on the use of membrane filters (0.22-|xm pore size) as a solid-phase matrix, which allows one to perform the washing steps under filtration.
Discussion Enzyme-linked immunosorbent assays are undoubtedly efficient tools in the diagnosis of infectious diseases, particularly of mycoplasmal infections. The development and the application of the pertinent techniques have been emphasized, and many laboratories have developed independent assays for their own purpose. Although the proposed ELISAs are all performed using the same basic procedures, many technical modifications have been described. This may lead to some confusion, especially for the novice, and reservation with regard to accepting the tests as representative techniques, especially since the majority of them are not properly validated. In the development of ELISAs, it is highly recommended to follow representative protocols for each assay, taking into account key conditions and an internal quality control with known reference standards. One of the major problems in connection with the indirect ELISAs used in the diagnosis of large animal mycoplasmosis is the preparation of suitable antigens. While the sonicated cells are mostly used as whole antigens, the specificity is very much enhanced by the selective solubilization of membrane proteins with a nonionic detergent (Tween 20). In a further step, chromatographic purification is very efficient, but may be cumbersome for certain laboratories. No specific antigens have been described so far for the mycoplasmas of the "mycoides group." The general situation points out the urgency of searching specific immunogenic and immunodominant antigens. However, this is not an easy task since a diagnostic test needs to detect not only persistent but also early produced antibodies and because the pathogenesis of large animal mycoplasmosis is not sufficiently elucidated for identifying specific virulence attributes. Because large animal mycoplasmosis is highly contagious and consequently of great socioeconomic importance, programs have been established for the control, surveillance, and eradication of these diseases in accordance with national or international schemes (International Office of Epizootics). Therefore, the development of standardized ELISA techniques offering high quality and good performance is imperative. Furthermore, the diagnostic validation of the technique is a prerequisite for general acceptation. Recommendations for standardization and validation have been formulated by an international expert group (Wright et al., 1993). The trend to international conformity should encourage all the research groups in this field, especially the industries that are increasingly interested in producing commercial kits, to devel-
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op and define new techniques more precisely and in accordance with fundamental rules. Because of the improved availability of monoclonal antibodies, the specificity of immunoassays can be increased considerably. These essays are, however, primarily indicated for use in the diagnosis of infections due to closely related mycoplasmas like the ones from the "mycoides cluster." Major technical problems still arise from the production, from the need of at least two pooled monoclonal antibodies for testing, and, if not commercially supplied, from poor availability to other laboratories. In conclusion, despite the wide use of ELISA techniques for the diagnosis of mycoplasmosis of large animals, a great deal remains to be done in order to improve the situation. A special effort is required in the standardization of the tests, particularly for those mycoplasmoses that are controlled internationally by official regulations.
References Brocchi, E., Gamba, D., Poumarat, F., Martel, J. L., and De Simone, F. (1993). Improvements in the diagnosis of contagious bovine pleuropneumonia through the use of monoclonal antibodies. Rev. Sci. Tech. Off. Int. Epizoot. 12, 559-570. Carpenter, A. B. (1992). Enzyme-linked immunosorbent assays. In "Manual of Clinical Immunology" (N. R. Rose et al., eds.), 4th ed., pp. 2-9. Am. Soc. Microbiol., Washington, DC. Guerin, C , Thiaucourt, F., Mady, V., Breard, A., and Lefevre, P. C. (1993). Rapid diagnosis of contagious caprine pleuropneumonia in pleural fluids by immunobinding assay. Small Rum. Res. 12, 193-200. Heller, M., Berthold, E., Pfiitzner, H., Leirer, R., and Sachse, K. (1993). Antigen capture ELISA using a monoclonal antibody for the detection of Mycoplasma bovis in milk. Vet. Microbiol, yi, 127-133. Kazama, S., Yagihashi, T., and Seto, K. (1989). Preparation of Mycoplasma hyopneumoniae antigen for the enzyme-linked immunosorbent assay. Can. J. Vet. Res. 53, 176-181. Nicolet, J., Paroz, P., and Bruggmann, S. (1980). Tween 20 soluble proteins of Mycoplasma hyopneumoniae as antigen for an enzyme-linked immunosorbent assay. Res. Vet. Sci. 29, 305309. Poumarat, F., Perrin, B., and Longchambon, D. (1991). Identification of ruminant mycoplasmas by dot immunobinding on membrane filtration (MF dot). Vet. Microbiol. 29, 329-338. Wright, P. F., Nilsson, E., Van Rooij, E. M. A., Lelenta, M., and Jeggo, M. H. (1993). Standardisation and validation of enzyme-linked immunosorbent assay techniques for the detection of antibody in infectious disease diagnosis. Rev. Sci. Tech. Off. Int. Epizoot. 12, 435-450.
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B4 ELISA IN HUMAN UROGENITAL INFECTIONS AND AIDS Richard Yuan-Hu Wang and Shyh-Ching Lo
Introduction Purpose
Identifying urogenital infections due to highly fastidious mycoplasmas, such as Mycoplasma genitalium, M. fermentans, or M. penetrans, is difficult in that culture is rarely successful and usual histopathological techniques do not stain the wall-free mycoplasmas in urine sediments. Polymerase chain reaction (PCR) of urines or urethral swabs may provide a highly sensitive assay to detect infections by these urogenital mycoplasmas (Dawson et al., 1993; Homer et al., 1993). However, special care is required to avoid contamination during the preparation of clinical samples for analysis. The complications have prevented most clinical microbiology laboratories from using PCR as a general tool for the diagnosis of mycoplasma infections. A sensitive and specific serological assay such as enzyme-linked immunosorbent assay (ELISA) or Western blotting for rapid detection of mycoplasmal infections, similar to the techniques used in many viral infections, is urgently needed. Successful development of a reliable ELISA to detect mycoplasma-specific antibodies will provide a powerful tool for clinicians and scientists to assist with clinical diagnoses and epidemiological studies.
115 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Rationale Since mycoplasmas have much greater antigenic complexity than viruses, the main challenge in the development of a meaningful serological assay is the ability to identify specific target antigens of mycoplasmas in the serological responses of human hosts. Mycoplasmal lipid-associated membrane proteins (LAMPs) are highly antigenic (Wang et al., 1992, 1993a; Wise et al, 1993) and are exposed on the cell surface (Wise, 1993). These attributes make them the most likely immunogenic targets for hosts' responses in mycoplasmal infections (Wang et al., 1993b). Most importantly, we have found that antibodies to LAMP antigens from each individual species of mycoplasma are highly species specific and are not cross-reactive with those from other species (Wang et al, 1992, 1993a,c). Serological assays using both ELISA and Western blots have been developed to detect specific antibodies to mycoplasmal LAMPs. These assays for M. penetrans, M. salivarium, and M. pirum have clearly demonstrated specificity and validity (Wang, et al., 1992). In subsequent studies, we further demonstrated the uniqueness of antigenic epitopes of LAMPs, even between those species of mycoplasmas previously known to be antigenically closely related, such as M. genitalium and M. pneumoniae (Wang et al, 1993c). Thus, we can develop an ELISA to detect mycoplasma-s^tcxfiQ antibodies using LAMPs prepared from each particular species of mycoplasma as target antigens.
Procedure Materials and Reagents Nunc-Immuno F96 MaxiSorp w/certificate plate (Nunc, Inc., Naperville, IL) and sealing tape for ELISA plate Bovine serum albumin (BSA) for overcoating ELISA plates (Cat. No. A3803, Sigma, St. Louis, MO) Normal goat serum (Life Technologies, Inc., Grand Island, NY) Phosphate-buffered saline (PBS) Nonidet P-40 (NP-40) (Sigma) Polypropylene conical tube (sterile) Eppendorf repeater pipettor and combitip Biotin-labeled antibody to human IgG(r) chain, peroxidase-labeled streptavidin, 2,2'-azinodi[3-ethylbenzthiazoline sulfonate] (ABTS) peroxidase substrate solution, and ABTS peroxidase stop solution (Kirkegaard & Perry Laboratories, Inc., Gaithersburg, MD)
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37°C incubator, cell sonicator, and ELISA plate reader equipped with automatic mixing and automatic subtraction of the reading of blank devices ELISA plate coating buffer: 20 mM sodium bicarbonate buffer, pH 9.6, 0.15 M NaCl Buffer A: 50 mM Tris-HCl (pH 8.0 at 25°C), 0.15 M NaCl, 1 mM EDTA Buffer B: 50 mM Tris-HCl (pH 8.0 at 25°C), 0.5 M NaCl 1 mM EDTA Buffer C: 50 mM Tris-HCl (pH 7.5 at 25°C), 0.25 M NaCl, 1 mM EDTA Buffer W: i x PBS, pH 7.0, 0.05% NP-40 10% Triton X-114 (v/v) in sterile deionized water and stored at 5°C Diluent I: 10% (v/v) normal goat serum, 2% BSA, 0.3% NP-40, 0.05% sodium azide, i x PBS, pH 7.2 Diluent II: 10% normal goat serum, 2% BSA, 0.3% NP-40, IX PBS, pH 7.2 Dilutent III: 10% normal goat serum, 2% BSA, 0.1% NP-40, IX PBS, pH 7.2
Methods ANTIGEN PREPARATION
1. Wash the mycoplasmas by resuspending the cell pellet from 2 liters of SP4 culture in 30 ml ice-cold buffer C. Spin at 20,840 g for 15 minutes at 5°C. Discard the supernatant. 2. Resuspend the pellet in ice-cold buffer A and adjust the protein concentration to 1 mg/ml. 3. Lyse the cells by adding Triton X-114 to 2% (v/v) from a 10% stock. Mix the lysates by vortexing. 4. Sonicate the lysates three times using a microtip (2 mm in diameter, 6 cm in length) submerged in the lysates. Each time consists of 45 seconds of sonication at 70% output power, followed by a 5-minute cooling period in ice. 5. Adjust the NaCl concentration to 0.5 M with a 5 M stock. Mix and keep the lysates in ice for 2 hours. Precool the centrifuge to 5°C for at least 20 minutes before carrying out the next step. 6. Centrifuge at 20,840 g for 20 minutes at 5°C to remove the insoluble materials. Transfer the supernatant to a new tube and store in ice. 7. Warm up the centrifuge to 30°C. 8. Start the first cycle of Triton X-114 phase fractionation by incubating the cleared lysates at 37°C in a water bath for 10 minutes to induce the condensation of Triton X-114. The lysates appear cloudy and partially biphasic. 9. Centrifuge at 5200 g for 10 minutes at 30°C. Some precipitates may appear in the bottom of tube, but need not be removed at this stage. 10. Remove the upper layer of aqueous solution as much as possible.
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11. Incubate the heavier detergent-enriched phase (5-10% of the original total volume) in ice. Add ice-cold buffer B up to the original volume. Mix thoroughly by vortexing and store in ice for 5 minutes. 12. Repeat step 8 through step 11 two more times. 13. After the third-cycle of Triton X-114 phase fractionation, resuspend the detergent-enriched phase in ice-cold buffer A up to the original volume. Mix by vortexing and store in ice for 30 minutes. 14. Centrifuge at 20,840 g for 20 minutes at 5°C. 15. Transfer the supernatant to a new tube and save as Triton X-114 extract. Make 0.8- to 1-ml aliquots in 1.5-ml sterile polyproplyene tubes and store at -70°C. The LAMP antigens are stable for serological assay under the storage conditions for at least 2 years. 16. Determine the protein concentration in the Triton X-114 extract by using the Bio-Rad (Hercules, CA) DC protein assay, a modified version of Lowry method, with bovine 7-globulin as standard. SERUM SAMPLE PREPARATION
1. Serum or plasma samples should be handled as potential infectious agents with precautions according to the NIH Biosafety guidelines. 2. Undiluted samples are stored at 5°C for short-term use (within a month); otherwise stored at -70°C. 3. Make a 250- to 500-|JL1 aliquot of 5- to 10-fold diluted solution for each serum sample in diluent I. Store these diluted samples at 5°C. ELISA
1. Thaw one tube of Triton X-114 extract at room temperature (25°C). Mix by vortexing and store in ice. 2. Make a 1:100 dilution of Triton X-114 extract in the ELISA plate coating buffer at 25°C in a 50-ml polypropylene conical tube. The concentration of protein is about 2 |xg/ml. 3. Coat the Nunc-Immuno F96 MaxiSorp plate with 100 |xl solution in each well using an Eppendorf repeater pipettor and sterile combitip. Note: We have found this type of ELISA plate most suitable due to the presence of detergent in the coating solution even at a very low concentration (<0.002%), the other kind of plate (PolySorp) from the same company has been found unworkable. 4. Cover the top of the ELISA plate with sealing tape, place the plate in a plastic box with a sheet of pre wetted gel blotting paper on the bottom, and incubate at 37°C for 4 hours. 5. Aspirate the coating solution and wash the plate twice with buffer W.
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Invert the plate and tap on two sheets of absorbent paper to remove residual fluids. 6. Overcoat the plate by adding 200 JJLI of 0.1% BSA and 0.02% sodium azide to each well and incubate at room temperature (25°C) for 2 hours. 7. Repeat step 5. At this stage, the ELISA plate can either be stored at -70°C for as long as 6 months or be processed for the next step. (For storage, seal the top of the plate with tape and then wrap in a plastic bag.) 8. Pipette 100 |xl of diluent II to each well, and then add 2-10 |JL1 of each 5 x diluted serum sample, depending on the final tested dilution. The first well on the plate should be set aside for a blank without adding serum sample. 9. Rock the plate on a orbit shaker at room temperature for 3 minutes. Incubate at 5°C overnight and then at 37°C for 2 hours. 10. Wash the plate six times as in step 5. 11. Prepare 1:1000 diluted biotin-labeled goat anti-human IgG(r) (0.5-mg/ml stock in 50% glycerol stored at -20°C) in dilutent III. Add 100 JJLI of this solution to each well and incubate at 37°C for 90 minutes. 12. Wash the plate as described earlier. Add 100 |xl of 1:20,000 diluted peroxidase-labeled streptavidin (0.5-mg/ml stock) in diluent III to each well and incubate at 37°C for 90 minutes. Note: There are great differences in terms of enzyme activities for peroxidase-labeled streptavidin from different vendors. It is necessary to titrate each batch of enzyme to find the appropriate dilution. 13. Wash the plate six times as in step 5. Prepare a 1:1 (v/v) mixture of hydrogen peroxide solution and ABTS substrate solution. Warm up the mixture at 37°C for 10 minutes. 14. Add 100 |xl of the ABTS-H2O2 mixture to each well. Develop the color reaction at 37°C for 20 minutes. 15. Stop the reaction by adding 100 jxl of 1% SDS (ABTS stop solution) to each well. 16. Measure the optical density (OD) of each well at 405 nm corrected with a reference wavelength at 650 nm and subtract the OD reading from the blank.
Quality Control / . Positive Control
Select a serum sample from patients infected by the mycoplasma as proven by culture or PCR. Make serial dilutions of the sample starting from 1:250 and find the end point of dilution which still has an OD reading equal to or greater than 1.5 in the ELISA test. Use this diluted serum sample as a positive control. In each run of ELISA tests, the OD value of the positive control should not be less
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Richard Yuan-Hu Wang and Shyh-Ching Lo
than 1.2 to consider the tests to be valid. Prepare the positive control in diluent I. Keep one aliquot of this diluted positive control serum at 5°C for daily use within a month and store the rest at -70°C. 2. Negative Control Include three serum samples from healthy blood donors which have OD values of less than 0.1 at the 1:250 dilution as negative controls in each run of the ELISA tests. The mean reading of these controls is included in the determination of the ELISA cutoff value in each test. 3. Criteria for Positive ELISA Test The original ELISA cutoff value is derived from the mean reading of more than 300 HIV (human immunodeficiency virus)-negative healthy blood donors plus four standard deviations. To correct the variation of readings in each test, the modified cutoff value is determined by the sum of the mean reading of three preselected negative controls and the adjusted original cutoff value, which is 0.25 for M. penetrans, 0.4 for M. fermentans, and 0.5 for M. genitalium. Serum samples with OD readings greater than the modified cutoff value are considered to be positive for the specific antibodies tested. All positive serum samples should be repeated twice and also tested in plates coated with BSA without mycoplasmal LAMP antigens to assess potential nonspecific binding problems of IgG in the serum samples.
Discussion It may be necessary to optimize the pH and the salt concentrations of the buffers used during the isolation of LAMP antigens by Triton X-114 fractionation for each individual species of mycoplasma tested. Sodium bicarbonate buffer, pH 9.6, plus 0.15 M NaCl is found to be the most effective in coating the LAMP antigens prepared by the just-mentioned technique onto ELISA plates. ELISA using LAMPs as target antigens prepared from different species of mycoplasmas provides a highly sensitive test for antibodies to the specific species of mycoplasma. The test is particularly helpful in studying unusual human mycoplasmas such as M. penetrans or M. genitalium. Our preliminary data showed that these hard-to-culture human urogenital mycoplasmas are transmitted primarily through various sexual contacts (Wang et al, 1992, 1993a,c). The sensitive assay may be less useful in studying the highly prevalent human mycoplasmas, such as M. salivarium, M. orale, or even M. pneumoniae, because a
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majority of individuals already test positive for antibodies to these mycoplasmas. At present, we still do not know if the difference of antibody titers to these mycoplasmas makes any difference in clinical considerations. The LAMPs prepared as described earlier are also suitable for Western blot analysis. In our laboratory, we routinely confirm the antibody reactivity to mycoplasmal LAMPs by Western blot (Wang et al, 1992, 1993c). Only those serum samples which test positive by ELISA repeatedly and produce positive reaction bands with the marker LAMPs of mycoplasma on Western blots are diagnosed as positive. Our preliminary experience showed that the ELISA could also be applied to detect specific IgA to these urogenital mycoplasmas with high sensitivity. The organisms apparently colonize the mucosal surface of the urogenital tract. Many patients who test positive for IgG antibodies to the mycoplasmas also test positive for IgA antibodies (Lo, S.-C, unpublished data). The application for detecting specific IgM to these mycoplasmas is currently being investigated.
References Dawson, M. S., Hayes, M. M., Wang, R. Y.-H. Armstrong, D., Budzko, D. B., Kundsin, R. B., and Lo, S.-C. (1993). Detection and isolation of Mycoplasma fermentans from urine of HIV positive patients with AIDS. Arch. Pathol. Lab. Med. 117, 511-514. Homer, P. J., Gilroy, C. B., Thomas, B. J., Naidoo, R. O. M., and Taylor-Robinson, D. (1993). Association oiMycoplasma genitalium with acute non-gonococcal urethritis. Lancet 342, 582585. Wang, R. Y.-H., Shih, J. W. K., Grandinetti, T., Pierce, P. F., Hayes, M. M., Wear, D. J., Alter, H. J., and Lo, S.-C. (1992). High frequency of antibodies to Mycoplasma penetrans in HIVinfected patients. Lancet 340, 1312-1316. Wang, R. Y.-H., Shih, J. W.-K., Weiss, S. H., Grandinetti, T., Pierce, P. F., Lang, M., Alter, H. J., Wear, D. J., Davies, C. L., Mayur, R. K., and Lo, S.-C. (1993a) Mycoplasma penetrans infection in male homosexuals with AIDS: High seroprevalence and association with Kaposi's sarcoma. Clin. Infect. Dis. 17, 724-729. Wang, R.-Y., Grandinetti, T., Melcher, G., Hendrix, C , Hayes, M. M., D. J., Shih, J. W.-K., and Lo, S.-C. (1993b). Seroconversion following Mycoplasma penetrans infection in patients with AIDS. Abstr. 93rd Gen. Meet. Am. Soc. Microbiol., G-17, p. 165. Wang, R.-Y., Grandinetti, T., Tully, J., Davies, C. L., Wear, D. J., Shih, J. W.-K., and Lo, S.-C. (1993c). Antibodies to M. genitalium in patients with AIDS and patients attending sexual transmitted diseases clinics. Abstr. 93rd Gen. Meet. Am. Soc. Microbiol. G-18, p. 165. Wise, K. S. (1993). Adaptive surface variation in mycoplasmas. Trends Microbiol. 1, 59-63. Wise, K. S., Kim, M. F., Theiss, P. M., and Lo, S.-C. (1993). A family of strain-variant surface lipoproteins of Mycoplasma fermentans. Infect. Immun. 61, 3327-3333.
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B5 ELISA IN RESPIRATORY INFECTIONS OF HUMANS Gail H. Cassell; Ginger Gambill^ and Lynn Duffy
General Introduction Ureaplasma urealyticum is a cause of pneumonia in newborn infants and is associated with chronic lung disease in very low birth weight infants. Mycoplasma hominis also causes pneumonia in newborn infants and has occasionally been associated with pneumonia in immunocompromised patients (Cassell et al., 1994). M. genitalium and M. fermentans have been associated with lower respiratory disease in adults, but their clinical significance in respiratory disease has not been systematically investigated. M. pneumoniae is a significant cause of respiratory disease in all age groups (Foy, 1993). Detection of specific antibody by the enzyme-linked immunosorbent assay (ELISA) is a reliable and useful method of diagnosis of M. pneumoniae respiratory disease in humans. Although ELISAs have been established for U. urealyticum (Brown et al., 1983) and M. hominis (Brown et al., 1987), these tests have not been evaluated for diagnosis of respiratory disease and are not discussed in this chapter. Diagnosis of M. pneumoniae Infection Approximately 10 to 20% of all cases of pneumonia are due to M. pneumoniae. In addition, this organism is a common cause of tracheobronchitis and other respiratory syndromes such as bronchiolitis and pharyngitis. Less common manifestations include croup, conjunctivitis, and otitis (Cassell et al., 1994). 123 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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Severe and extensive pulmonary disease occurs occasionally in M. pneumoniae infections. Extrapulmonary manifestations (dermatologic, neurologic, cardiac, and renal) occur and in some cases patients have no prior respiratory manifestations. The diverse clinical illnesses associated with M. pneumoniae infection may lead to a delay in definitive diagnosis. The clinical manifestations and radiographic and laboratory findings in M. pneumoniae infection are not distinctive enough to allow an accurate diagnosis. Definitive diagnosis should include cultural isolation of M. pneumoniae from bronchial secretions or other sites in the respiratory tract; isolation of the organism when appropriate from extrapulmonary sites; and a fourfold rise or fall in specific antibody (an increase 3 to 9 days after onset of symptoms and a peak at 3 to 4 weeks). A single high titer is suggestive but not definitive. Cold agglutinins develop in approximately 50% of individuals infected with M. pneumoniae. Cold agglutinins appear during the second week of illness, disappear in 3 to 6 months, but they are nonspecific and can develop in patients with other diseases, including viral and bacterial respiratory infections. Cultural isolation of M. pneumoniae and positive identification may require 3 to 4 weeks. This slow growth coupled with the lack of suitable commercially available culture media limits the usefulness of cultural diagnosis and results in a reliance on serology alone or serology and one of the newer techniques like an ELISA-based antigen detection assay or polymerase chain reaction (PCR). However, the latter have not been extensively applied and rigorously compared to the gold standard of culture in large numbers of patients. Because M. pneumoniae may persist for several months following acute infection, both antigen detection and PCR require confirmation by assessment of the serological response to verify that the infection is current and that positive results are not the result of continuing carriage of the organism from a previous infection, unrelated to the current episode under investigation (Marmion et al., 1993). Thus reliable serology is critical for accurate diagnosis of M. pneumoniae respiratory disease. The ELISA for detection of antibody appears to be the method of choice. Comparison of ELISA to Other Serological Diagnostic Methods for M. pneumoniae Infection The specific antibody response to M. pneumoniae infection may be demonstrated by indirect hemagglutination, growth inhibition, immunofluorescence, complement fixation, or ELISA. Growth-inhibiting antibodies are specific, rise within a few weeks after infection, peak at approximately 4 months, and persist for long periods. Complement-fixing (CF) antibodies are the most widely used for serodiag-
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nosis of M. pneumoniae infection (Kenny, 1992). CF antibodies rise rapidly, peak at 1 month after infection, and persist for a shorter time, but their analysis is satisfactory for routine diagnostic purposes. In fact, in an extensive comparison of CF with culture in 674 patients the sensitivity and specificity of the CF test was shown to be 90 and 94%, respectively (Kenny et ai, 1990). In contrast, the sensitivity of culture was only 64% for persons with serological evidence of infection. In this 12-year study, the geometric mean titer was 1:2.8 in the acutephase sera which was the same as that of culture-negative individuals. This indicates that the diagnosis of acute infection should not be limited to the CF test. Furthermore, since the CF test measures predominantly "early" IgM antibodies to M. pneumoniae and only to a minor extent IgG antibodies, the diagnostic value of the CF test may be limited to initial M. pneumoniae infection (Jacobs, 1993). This can be problematic because of the rather slow onset of clinical symptoms and the failure of patients to seek early treatment. The ELISA offers several major advantages over other antibody detection methods: objectivity, immunoglobulin class and subclass specificity, and increased sensitivity. Sample volumes necessary for assay are small (10-100 |xl). We have compared the efficacy of an IgM and IgG ELISA based on whole organisms of the FH reference strain as the antigen, to that of culture and PCR for the diagnosis of M. pneumoniae infection in 256 children (2 to 16 years of age, mean 6.3 years) with radiographically confirmed pneumonia. Of 38 who had all three diagnostic tests performed and which were positive by one or more tests, 35 were positive by ELISA and 32 of these were positive by culture, PCR, or both. Fourteen of the 16 culturally positive patients were also positive by ELISA. Thus this ELISA appears more sensitive than culture and comparable in sensitivity to detection of infection by PCR. The fact that only 3 of the 35 ELISA positives were negative by the other methods also suggests that this ELISA is highly specific.
Antigen Preparation Introduction M. pneumoniae ELISA antigens which have been used and shown to be acceptable include: washed whole organisms, sonicated whole cell antigen preparations, detergent-lysed organisms, and microtiter plate-cultured and formalinfixed mycoplasmas. The antigen preparation must be chosen at the discretion of the individual investigator and tailored to the desired uses of ELISA. The method for whole cell antigen is given in detail here.
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Materials 250-ml sterile disposable tissue culture flasks 3TC incubator and 4°C refrigerator Logarithmic broth culture of M. pneumoniae (1-3 liters) grown in SP4 broth (see Chapter A2 in Vol. I). Phosphate-buffered saline containing thimerosal, sterile (PBS/M) Centrifuge bottles (250 ml), sterile High-speed, refrigerated centrifuge Blood agar plate for the monitoring of sterility Spectrophotometer pH meter
Formula for PBS/M (Phosphate-Buffered Saline, pH 7.2 to 7.4, 0.01 M Phosphate and 0.15 M NaCI with Thimerosal) 1. For 4 liters of 10 x PBS/M: Dissolve reagents in 3 liters of distilled water and make up to 4 liters. Store at room temperature. NaH2P04-H20 (sodium phosphate monobasic) 10.25 g Na2HP04 (sodium phosphate dibasic) 47.75 g NaCl 350.65 g Thimerosal 4 g
2. For 1X PBS/M, dilute 10 x stock 1:10 (final buffer of 0.01 M phosphate and 0.15 M NaCl); pH solution to approximately 7.33. Store at room temperature.
Procedure 1. Dispense 125 ml SP4 broth into each of four sterile tissue culture flasks and add 1 ml FH strain of M. pneumoniae reference stock (American Type Culture Collection, No. 15531). 2. Incubate at 37°C until pH-induced color changes from red to orange/yellow (about 3 to 4 days). Wash cells by decanting medium from flask into 10% Clorox disinfectant. Add 5 ml of PBS/M to each flask and wash the monolayer of cells, discarding fluid also into disinfectant. Repeat wash two times. 3. Add 5 ml of PBS/M to each flask and scrape the sides and bottom of the flask with a tissue scraper to remove attached organisms. Pool organisms from each flask and centrifuge at 10,000 g, 4°C, for 20 minutes. 4. Wash pellet of organisms with 5 ml PBS/M and centrifuge again at 10,000 g for 20 minutes. Resuspend organisms in 5 ml of PBS/M and dilute this
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suspension to an optical density (OD) reading equal to 0.10 at 600 nm. Store at 2°C-8°C in PBS/M. 5. Plate an aliquot (0.025-0.05 ml) of the cell suspension on blood agar and incubate for 72 hours at 37°C to ensure that no bacterial contamination has occurred. 6. This stock solution should be further diluted for coating of plates (see Establishment and Performance of the Assay).
Discussion
The influence of antigenic heterogeneity of M. pneumoniae on serologic diagnosis using any method has not been rigorously evaluated. However, immunoblotting studies indicate that the principal protein antigens are stable in strains collected over a 10-year period. Large multicenter trials indicate that those individuals who are culture positive are almost always serologically positive. This also suggests that strain variation may not have a major impact on serological diagnosis. However, this should be examined more systematically.
Conjugate Introduction
The ability to detect a class-specific antibody is useful for evaluating the immune response in individuals infected with M. pneumoniae. High quality, conjugated, class-specific anti-immunoglobulins are commercially available. Materials
Alkaline phosphatase-conjugated anti-human IgG (FC) (Jackson Immuno Research Labs, Inc., Cat. No. 109-055-098 or equivalent) Alkaline phosphatase-conjugated anti-human IgM Fc5u fragment specific (Jackson Immuno Research Labs, Inc., Cat. No. 109-055-129) 10% fetal calf serum (PCS) (PCS purchased from Hyclone Laboratories, Logan, UT). Dilute 10 ml/100ml IX PBS/M. Make fresh for each experiment.
Procedure
1. Reconstitute reagents and store according to package insert. Pay particular attention to expiration dates.
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2. Dilute to a proper working dilution in 10% FCS. The working dilution used for each lot should be determined by titration. Note: The working dilution should be prepared fresh for each experiment. The proper diluent may vary depending on the manufacturer of the conjugate. Normal horse serum in PBS/M (10%) or 10% normal goat serum/PBS/M may be used with some conjugates.
Substrate Materials /7-Nitrophenyl phosphate (PNPP) (Sigma 104 phosphatase substrate) 5-mg tablet, Sigma Cat. No. 104-105 or equivalent. Light sensitive. Store at -20°C in the dark.
Procedure Prepare working substrate during incubation of ELISA plates with labeled antibody. The volume of substrate to be prepared must be determined for each run (one tablet/5 ml). Each well uses 0.1 ml. The final concentration is 1.0 mg PNPP/ml diluent. Wrap in tin foil or otherwise keep in the dark as material is light senstive.
Establishment and Performance of the Assay Equipment and Materials Micro ELISA autoreader (MR-580 by Dynatech, Flow Titertek Multiskan, or equivalent) Single and multichannel pipettes and appropriate tips that will deliver 2, 50, 100, 300, and 1000 |xl Microtitration reservoir (Flow Labs 77-824-01 or equivalent) Manifold plate washer Water bath capable of maintaining 45° 2°C 96-well EIA plates (ICN-Flow, Cat. No. 76-381-04, or equivalent) Adhesive tape for sealing EIA plates 10-ml serological pipettes 12 X 75-mm polypropylene tubes for serum dilution Antigen
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Conjugate Substrate Formulas for
Buffers
A. 1 % BSA/PBS
1 g bovine serum albumin (Fraction V)/100 ml PBS/M Make fresh for each experiment. B. PBS/M TWEEN 20 (0.5% TWEEN)
10 ml Tween 20 per 2 liters of 1 x PBS/M. Store at room temperature. C. 1 N NaOH
40 g NaOH per liter of distilled H2O. Store at room temperature. D. SUBSTRATE BUFFER
0.05 M Na2C03 buffer with O.OOI M MgCl2 at a pH of 9.8 Solution 1: 5.2995 g Na2C03/liter of distilled H2O Solution 2: 4.2005 g NaHC03/liter of distilled H2O Solution 3: 20.330 g MgCl2-6H20/liter of distilled H2O Make 1 liter of all three solutions. To each liter of solutions 1 and 2, add 10 ml of solution 3. Add solution 1 to solution 2 until a pH of 9.8 is achieved. Solution 3 may be stored at room temperature for future use. Store the prepared buffer at 4°C. E. ANTIGEN BUFFER
Solution 1: 8.401 g NaHC03/liter of distilled H2O Solution 2: 10.599 g Na2C03/liter of distilled H2O Make one liter of each solution. To each solution add 0.2 g NaN3 (sodium azide). Add solution 2 to solution 1 until a pH of 9.6 is reached. Store at 4°C. When properly stored, the antigen buffer is stable indefinitely. After use, the antigen buffer should be discarded in a biohazard bag for incineration because it contains NaN3. F. WATER REQUIREMENTS
Milli-Q deionized water is needed for the preparation of PBS/M, antigen buffer, substrate buffer, and 1 A^ NaOH.
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Collection and Handling of Patient Serum
Blood obtained by venipuncture, with no anticoagulant, should be allowed to clot and then centrifuged. The serum can be stored up to 2 days at 2°-8°C. For longer storage, freeze samples at — 20°C in a non-frost-free freezer or preferable at — 70°C, if available. Repeated freezing and thawing may cause deterioration of the test sample. A serum containing visible particulate matter should be clarified by low-speed centrifugation before testing. Heat inactivation is not necessary. Sample size is 0.5 ml of serum. Note: All serum from humans should be considered potentially infectious [e.g., hepatitis, acquired immunodeficiency virus (AIDS), or other agents] and handled accordingly. Negative Serum Pool (NSP)
A pool of 20 to 25 negative serum samples for use in each assay to determine the cutoff point between negative and positive samples is prepared by pooling 1.0-ml volumes of the negative samples. The samples chosen for the pool must give a balanced representation of OD readings for negative samples. Each new negative pool must be extensively evaluated before being used. The pool must be evaluated using at least 50 normal individuals to determine cutoff values (usually 3 SD above the mean of the 50 normal sera). Following standardization of the NSP, freeze in aliquots. Working NSP is prepared by making a fresh 1:1000 dilution in 1% BSA, fresh for each experiment. Procedure DETERMINATION OF OPTIMUM ANTIGEN AND CONJUGATE DILUTIONS
1. To determine the optimum antigen dilution, test various dilutions of antigen (twofold dilutions in antigen buffer starting at 1:20) with a high concentration of known immune serum and a high concentration of conjugate. Thirty minutes prior to the time serum is added, decant antigen and add 0.3 ml/well of 1% BSA/PBS/M to block unbound sites. 2. Plot the antigen concentration against absorbance at 405 nm which will give a sigmoidal curve. The highest dilution of antigen that is still on the upper plateau is the dilution of antigen (or perhaps twice that concentration) to use in the assay as it ensures antigen excess without antigen waste. 3. To determine the optimum conjugate dilution, incubate different conjugate dilutions in 10% FCS/PBS/M (dilutions of conjugate to test initially are 1:250, 1:500, 1:1000, 1:2000, 1:3000, 1:4000, and 1:5000) with the set antigen dilution determined as described earlier. Plot the conjugate dilution against the absor-
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bance at 405 nm and a sigmoidal curve will be generated. Select the conjugate dilution for routine use that is the lowest dilution (highest concentration) which is on the lower plateau as this ensures conjugate excess with a minimum of nonspecific binding. 4. The optimum conditions for ELISA are those that give a difference of >0.2 OD between hyperimmune and normal serum. If the normal serum continues to give high absorbency, this may be attributed to nonspecific binding.
Worksheet Preparation Daily worksheets should include lot numbers of antigen, substrate, conjugate, and EI A plates; location of controls and patient specimens on plate; control and patient results; dates of coating and blocking of plate; testing date; technician's name; and working dilutions of serum, antigen, and conjugate. Printout tapes from the EIA reader should be labeled and stapled to the worksheet for proper documentation. Worksheets should be filled out in duplicate, one for each subclass of antibody to be tested. Note: Patient samples and controls are to be done in duplicate. At least six negative serum pools are tested per plate for IgG and IgM. EIA Plate Preparation 1. For each plate to be coated, prepare 7-8 ml of a previously designated working dilution of M. pneumoniae antigen in antigen buffer. The optimum antigen dilution must be determined for each new lot by titration. Use the same lot number of plates for each run. 2. Using a multichannel pipette, add 100 |JL1 of diluted antigen to the inside 60 wells of the microtiter plate. Note: Wells in the outside of the EIA plates are not used due to uneven heating. Add 100 JJLI of PBS/M to all outside wells. 3. To complete the coating process, cover the plates with tape and store at 2°-8°C overnight. It is preferable to coat only enough for the next day's experiment. 4. When ready to use plate, decant antigen coating liquid into a biohazard bag, and tap plate vigorously on several thicknesses of disposable absorbent towel to drain the plate. 5. Using a multichannel pipette, add 300 |xl/well of 1% BSA-PBS/M to all 60 inside wells. Incubate for at least 30 minutes at 15° to 30°C. This blocks remaining binding sites on the plastic surface of the wells and decreases background interference.
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Preparation of NSP and Positive Reference Serums
1. While plate is blocking, prepare a 1:1000 dilution of negative serum pool (NSP IgG and IgM) and also dilute patient serum (0.002 ml serum + 2.0 ml 1% BSA/PBS/M). Mix well. Label tube 1:10 dilution. 2. For all positive control serums, prepare 1.0-ml volumes of twofold serial dilutions using 1% BSA/PBS/M.
Performance of Assay
1. Discard blocking reagent in sink and tap plate vigorously on several thicknesses of disposable absorbent towels. Add 100 jxl of prediluted negative serum pool and positive control samples to appropriate wells following daily worksheet. Also, add 100 |JL1 of 1% BSA/PBS/M to three wells for background control check. 2. Transfer 100 |xl of patient serum dilution to appropriate wells. Cover the plate with tape and incubate in a water bath at 45° 2°C for 30 minutes. 3. Discard the contents of wells in a biohazard bag. Fill wells completely with PBS/M Tween wash solution (0.5% Tween) using a 12-pronged manifold EIA plate washer attached to a 1-liter gravity wash bottle. Aspirate. Shake out excess washing fluid into a biohazard bag. 4. Repeat step 3 three additional times. 5. Empty the EIA plate, tap vigorously on several thicknesses of absorbent disposable towels, and add 100 |xl of diluted IgG antibody to all IgG plates and 100 |xl of diluted IgM antibody to all IgM plates. Seal with tape and incubate for 30 minutes at 45° 2°C. 6. Repeat steps 3 and 4. Add 100 |xl of substrate to each well. Add 100 |JL1 of substrate to one row of a blank plate to use in the ELISA reader. Seal with tape and incubate at 45°C ( 2°) for 30 minutes. 7. Stop reaction by adding 50 yA of I N NaOH to each well. 8. Take a blank EIA sample reading with a 405-nm filter. Thoroughly wipe dry the bottom of the ELIS A blank and test plates, being careful not to scratch the bottom of the plate. Determine OD units of each well.
Calculations
1. Determine mean OD values of 1% BSA diluent control. Subtract this value from the OD value of each NSP, positive control, and patient sample. 2. Determine the mean OD value of the patient's duplicate wells. Also determine the mean OD value of the six NSP wells on each plate.
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3. Multiply the mean OD values of the normal serum pool times 3.5 (3 SD) above the mean of 50 normals to determine the cutoff for the IgM antibody. 4. Multiply the mean OD value of the normal serum pool times 2.8 (3 SD) above the mean of 50 normals to determine the cutofffor the IgG antibody. Note: Perform steps 1-4 for each plate.
Interpretation of Results 1. The OD readings of any patient dilutions that are equal to or greater than the cutoff value are considered positive. Any patient sample that has a positive value in dilution 10 must be repeated and titered with twofold dilutions to an end point. The last dilution that is positive is the titer. Note: If one OD value on a sample is greater than the cutoff and the other OD value on the duplicate well is less than the cutoff, then that sample should be repeated. 2. Any sample with OD readings less than the cutoff value is negative. Patient samples negative in dilution 10 are reported as less than 10 (this is the range of normal serum). 3. Report titers for both IgM and IgG.
Clean-Up Procedure Discard all materials that have come in contact with biological material in a biohazard bag to be incinerated. Paper products, boxes, and other materials that have not come in contact with biological materials should be discarded in regular trash. Quality Control Procedures 1. Controls to be included in each assay, (a) Conjugate control: Antigencoated well in which PBS/M has been substituted for serum, (b) Substrate control: Antigen-coated well in which PBS/M is substituted for both serum and conjugate, (c) Reference sera: Immune and normal serum standards allow immediate recognition of deterioration of reagents. A previous patient positive for both IgG and IgM antibodies should be titered in each run starting with one dilution below the established end point and titering through a fourfold dilution above the established titer. The six normal serum pool wells on each plate are used as a negative control. 2. Determination of assay validity: Intraassay variability is generally a reflection of error in technique. If the controls fall outside 1 dilution of a previously determined titer, the run is considered invalid and must be repeated. The ELISA
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is a sensitive technique; care should be taken in pipetting and handling all reagents. Pipetting and mixing errors will be magnified. Cross-contamination between reagents, such as enzyme with substrate, will invalidate the test. Reusable glassware must be washed and thoroughly rinsed free of all detergent. Sterile disposable glass- or plasticware is preferred if proper rinsing cannot be assured. 3. Parallel testing: (a) when changing to a new lot number of antigen or conjugate, titrations must be done to determine the correct working dilution to use. (b) When a new negative serum pool is prepared, extensive parallel testing is required to maintain the normal values as already established. The OD units should be similar to the present NSP. For this reason, it is advisable to prepare a pool with a minimum volume of 20 ml. (c) New lot numbers of EIA plates should be checked on receipt in parallel by performing a duplicate assay along with a run performed on the present lot number of plates. Results should be similar before new plates are used for patient runs, (d) To calibrate pipettes, follow manufacturer's suggestions for schedule and method. The optics of the Flow Titertek Multiscan are to be checked according to manufcturer's suggestions, (e) When determining antigen and conjugate working dilutions, use all of the same plate, substrate, conjugate, etc., lot numbers that will be used in the actual test.
Discussion Antigen Specificity One of the major limitations of serological diagnosis of M. pneumoniae infection by any method is the antigenic cross-reactivity with M. genitalium. Although M. genitalium was originally isolated from the urogenital tract, it has occasionally been detected in the respiratory tract of patients with pneumonia. Confirmation of ELISA positives by immunoblotting of patient sera with M. pneumoniae and M. genitalium can minimize problems of cross-reactivity. Alternatively, short synthetic peptides deduced from the PI adhesin of M. pneumoniae and the MgPa adhesin of M. genitalium have been used to determine major common and species-specific epitopes in the ELISA. Peptides from the nonhomologous regions appear to overcome the problem of cross-reactions but ELISAs using these synthetic peptides have not been systematically evaluated using large numbers of patient specimens. The good agreement between the ELISA and M. pneumoniae-specific immunoblotting, PCR, and culture argues that cross-reactivity with M. genitalium, at least in some patient populations, is not a major problem. In diagnosing respiratory disease in patients at high risk for sexually transmitted diseases this may not be true.
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Isotype Determination and Impact of Patient Age in Diagnosis of M. pneumoniae Infection
Specific M. pneumoniae antibodies of the M, G, A, and E immunoglobulin classes have been detected by ELISA. However, the temporal sequence of antibody formation and persistence has not been systematically evaluated. Regardless, a number of commercially available M. pneumoniae diagnostic kits as well as diagnostic laboratories use tests which will only detect IgM antibodies. This is inappropriate based on current evidence and can result in misdiagnosis in over 50% of cases, depending on the patient population. Children and teenagers respond predominantly with IgM antibodies, whereas patients older than 40 years often have an IgG (56% of cases) response only, probably because of reinfection (Uldum et al., 1992). We have also compared diagnostic methods in children 2 to 12 years of age and in adults. Only 3% of children had an IgG response compared to 50% of adults who had only an IgG response. Thus had we used an assay which detected only IgM, we would have missed 50% of adult patients, most of whom were also culture and/or PCR positive and with radiographically confirmed pneumonia. Specific IgA antibodies against M. pneumoniae develop more regularly and more rapidly than IgM in adult patients with pneumonia (Granstrom et al., 1994). However, IgA titers also start to decrease earlier than IgM or IgG. For detection of IgM antibody, the same antigen can be directly conjugated to alkaline phosphatase and used in a J|L capture format (Granstrom et al., 1994). Increasing M. pneumoniae infection and disease in children and infants from 0 to 5 years are being reported. The influence of age on the antibody response has not been studied extensively. However, in at least two studies of infants of less than 12 months in which disease was diagnosed radiologically and the organism was isolated from throat cultures, there was no antibody response. This was in great contrast to preschool and school children in which disease and cultural isolation resulted in significant antibody responses. Since M. pneumoniae infection is often diagnosed only by serology, the failure of infants to mount an antibody response may have led erroneously to the belief that this organism does not cause disease in this age group.
References Brown, M. B., Cassell, G. H., Taylor-Robinson, D., and Shepard, M. C. (1983). Measurement of antibody to Ureaplasma urealyticum by an enzyme-linked immunosorbant assay and detection of antibody responses in patients with nongonococcal urethritis. J. Clin. Microbiol. 17, 288295. Brown, M. B., Cassell,G. H., McCormack, W. M., and Davis, J. K. (1987). Measurement of antibody to Mycoplasma hominis by an enzyme-linked immunosorbant assay and detection of antibody responses in women with postpartum fever. Am. J. Obstet. Gynecol. 156, 701-708.
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Cassell, G. H., Waites, K. B., and Crouse, D. T. (1994).Mycoplasmal infections. In "Infectious Diseases of the Fetus and Newborn Infant" (J. S. Remmington and J. O. Klein, eds.), pp. 619656. Saunders, Philadelphia. Foy, H. M. (1993). Infections caused by Mycoplasma pneumoniae and possible carrier state in different populations of patients. Clin. Infect. Dis. 17(Suppl. 1), S37-S46. Granstrom, M., Holme, T., Sjogren, A. M., Ortqvist, A., and Kalin, M. (1994). The role of IgA determination by ELISA in the early serodiagnosis of Mycoplasma pneumoniae infection, in relation to IgG and mu-capture IgM methods. J. Med. Microbiol. 40, 288-292. Jacobs, E. (1993). Serological diagnosis in Mycoplasma pneumoniae infections: A critical review of current procedures. Clin. Infect. Dis. 17(Suppl.), S79-S82. Kenny, G. E. (1992). Serodiagnosis. In "Mycoplasmas: Molecular Biology and Pathogenesis" (J. Maniloff, R. N. McElhaney, L. R. Finch, and J. B. Baseman, eds.), pp. 506-512. Am. Soc. Microbiol., Washington, DC. Kenny, G. E., Kaiser, G. G., Cooney, M. K., and Foy, H. M. (1990). Diagnosis of Mycoplasma pneumoniae pneumonia: Sensitivities and specificities of serology with lipid antigen and isolation of the organism on soy peptone medium for identification of infections. J. Clin. Microbiol. 28, 2087-2093. Marmion, B. P., Williamson, J., Worswick, D. A., Kok, T. W., and Harris, R. J. (1993). Experience with newer techniques for the laboratory detection of Mycoplasma pneumoniae infection. Clin. Infect. Dis. 17(Suppl. 1), S90-S99. Uldum, S. A., Jensen, J. S., Sondergard-Andersen, J., and Lind, K. (1992). Enzyme immunoassay for detection of immunoglobulin M (IgM) and IgG antibodies to Mycoplasma pneumoniae. J. Clin. Microbiol. 30, 1198-1204.
B6 MONOCLONAL ANTIBODIES AS DIAGNOSTIC TOOLS Chester B. Thomas^ Monique Gamier, and John T. Boothby
General Introduction A persistent theme in diagnostic mycoplasmology continues to be the need for rapid, definitive methods to detect either the presence of mycoplasmas in clinical specimens or an immune response to them. Immunoassay methods which determine the presence and antigenic identity of agents are supplanting isolation as the method of choice for detection of many pathogens in medical, veterinary, and plant pathology diagnostic laboratories. Monoclonal antibody (MAb) reagents, which retain uniform characteristics over time and batch, and which provide improved immunologic specificity, have substantially aided this trend since the early 1980s. In mycoplasmology, where species-level taxa have been defined serologically, elimination of poly specificity in diagnostic reagents is the compelling advantage for use of MAbs in immunoassay. Detection of mycoplasmal infection using monoclonal antibodies has been successful in a number of formats (Ball et al, 1990; Fos et al., 1992; Morsy et al., 1992) (See also Chapter B3, this volume). Detection of specific antibodies in serum using monoclonal-mediated ELISA and metabolic inhibition can determine the specificity of responses with pinpoint (epitope) accuracy (Feldman et al., 1992; Sorensen et al., 1992). Development of an immunoassay employing MAbs requires thorough characterization of the specificity of the MAb reagent as a prerequisite. Choice of an immunoassay format should begin with the specification of the performance goals of the assay. Among the things to consider are 137 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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the sample capacity needs, maximum time to report results, cost and availability of materials and equipment, and thorough consideration of how the test results will be used. In the latter regard it is important to consider what further action may be taken as a result of a positive or a negative test report. Will confirmatory testing be undertaken? If not, what are the risks or costs associated with actions based on false-positive or false-negative tests? It is beyond the scope of this chapter to enter into a full discussion of the alternative immunoassay formats and their relative merits; similarly, strategies for the production and selection of hybridoma cell lines and production of MAb reagents will not be covered. However, three procedures which typify the application of MAbs in immunoassays for rapid detection of mycoplasmas are presented.
Detection of Mycoplasma bovis in Bovine Milk INTRODUCTION
M. bovis mastitis of the dairy cow is a contagious infection of the mammary gland that can be spread rapidly in a milking herd. Transmission is via the teat orifice by contact with milking equipment contaminated with the infected milk of herd mates. Treatment of mycoplasma mastitis with antibiotics has not been shown to be effective. Control of herd outbreaks is dependent on measures that improve milking hygiene, to prevent or slow the spread of new infections, and the identification and segregation or removal of infected animals. The basic unit of risk for new infections is each time that a cow is milked, which puts a premium on rapid availability of test results to detect infected cows. Milking herds are often large, several hundred to a thousand or more, which can complicate the logistics of testing and slow the availability of culture-based test results. Often new infections occur in the interval from the collection of milk samples to the time when test results become available which may necessitate additional testing. Because the outcome of a positive test is often sale of the animal to slaughter, with an attendant economic loss of several hundred dollars, there is also a premium on avoidance of false-positive test results. The following method is used for detecting M. bovis in milk by an immunoassay using monoclonal antibodies. The format of this assay is a liquid-phase competition ELISA (C-ELISA) in which an enzyme-labeled M. bovis-spccific MAb is mixed with a milk sample to allow binding with M. bovis cells, if they are present in the sample. Subsequently, unbound labeled MAb is detected by reaction with solidphase bound M. bovis antigen in microtiter assay. If M. bovis is present in the milk there is reduced color in the microtiter assay. The advantages of this C-ELIS A format are high sample capacity, rapid (4- to 6-hour) test results, and a high positive predictive value of positive test results. This assay is appropriate as a screening test for the rapid identification of acute M. bovis mastitis.
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Antigen Detection Competition Enzyme-Linked Immunosorbent Assay MATERIALS Equipment
Flat-bottom, 96-well microtiter plates (Coming) Pipettes (single channel, PIO, P200, and PIOOO; multichannel adjustable from 100 to 200 fjLl) ELISA reader at 405 nm Buffers
Coating buffer: Na2C03, 1.59 g; NaHC03, 2.93 g; NaN3, 0.2 g; H2O for 1 liter after adjustment of the pH at 9.6 Phosphate-buffered saline (PBS) buffer: NaCl, 8.0 g; KH2P04-12H20, 2.9 g; NaN3, 0.2 g; H2O for 1 liter after adjustment of the pH at 7.4 Washing buffer: PBS containing 0.05% Tween 20 (PBS-T). Citrate buffer: dissolve 10.6 g citric acid in distilled H2O, adjust pH to 4 with 1 A^ NaOH, and bring to 1 liter with distilled H2O. Reagents
MAb-HRP conjugate: purified M. Z7^v/5-specific MAb conjugated to horseradish peroxidase (HRP) diluted to a predetermined optimum concentration in PBS containing 0.005% Tween 20. Color reagent: 0.4 mM ABTS; dissolve 0.27435 g ABTS (2,2'-azino-di[3-ethylbenzthiazoline-6-sulfonate]) in 12.5 ml distilled H^O Enzyme substrate: 1.5 mM H2O2; 0.5 ml 30% H2O2 in 7.5 ml distilled H2O Substrate solution: For 100 ml combine 1.0 ml color reagent, 0.3 ml enzyme substrate, and 98.7 ml citrate buffer. Prepare fresh. Stop reagent: 0.005% NaN3; 0.00325 g NaN3 in 1 liter distilled H2O. PROCEDURES
1. Sensitize microtiter plates with M. bovis antigen; dispense 100 |xl per well of antigen solution (5 |xg protein of washed whole M. bovis cells per ml of coating buffer), incubate for 4 hours at 37°C or overnight at room temperature. 2. Mark a set of tubes (12 x 75mm) to match the unknown samples, the M. bovis antigen standards, the antigen control, and an ELISA blank. 3. Transfer 500 |xl of each unknown sample, standards, or antigen control (PBS only) to the appropriately marked tubes and add 500 |xl of MAb-HRP. Mix well and incubate with occasional agitation for 1 hour at room temperature.
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4. Prepare another tube to be used as an ELISA blank (500 |xl PBS). To this tube add 500 fxl of PBS-T. Incubate as described eariier. 5. Wash the M. bovis antigen-sensitized microtiter plates three times with wash buffer. 6. Transer 100 jxl from each tube (unknown samples, M. bovis antigen standards, antigen control, and an ELISA blank) to the appropriate wells of each plate. 7. Incubate plate(s) for 1 hour at room temperature. 8. Wash plates five times with wash buffer. 9. Add 100 |JL1 substrate solution to each well of the plates. After 30 minutes stop the reaction with the ELISA stop solution. 10. Using the ELISA reader determine the A405 (using the ELISA blank as A405 = 0) for each well. 11. Graph the results for the M. bovis antigen standards (A405 vs protein concentration) and determine the concentration of M. bovis antigen in the samples. Any proportionate decrease in color will represent the presence of M. bovis in the sample. (A standard curve can also be prepared comparing M. bovis protein content vs colony-forming units (CFU)/ml or color changing units (CCU)/ml.) DISCUSSION
The antigen detection C-ELISA method presented here has successfully overcome the traditional serological problems of specificity and reproducibility (Thomas et aL, 1987). The monoclonal antibody used in this assay has been shown to be monospecific forM. bovis (Boothby etal., 1986a). The immortality of the antibody avoids problems of reproducibility associated with polyclonal antiserum. The limitation to generalized use of this assay is its sensitivity. As presented here, we successfully detected less than 500 ng M. bovis protein. This represents about 10"^ CFU/ml in the assay as conducted on field samples, which is suitable for detection of acute M. bovis mastitis. However, in the chronic phase of infection the number of CPU of M. bovis shed in milk may be well below this level, requiring the use of standard culture methods to be detected (Boothby et aL, 1986b). Some improvement in sensitivity can be addressed by means of a capture ELISA such as the one presented in the next section (DASELISA). A similar method detected 10^ CFU/ml M. bovis in milk, a twofold log improvement in sensitivity over the C-ELISA presented here (Heller et aL, 1993). Such a solid-phase ELISA is an ideal system for use with monoclonal antibodies because of its enzymatic amplification and thus high sensitivity; but this sensitivity is not without potential loss of specificity. Any number of difficulties in performing a solid-phase ELISA can lead to the production of a nonspecific color reaction which will be interpreted as the presence of antigen.
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Heller et al. (1993) reported a 5% false-positive test rate which increased to nearly 10% when milk samples were preincubated for 48 hours. One advantage of the C-ELISA over such methods is that the presence of antigen is only indicated by the inhibition of color production. This robust character of competition assays will make them more appealing whenever there is a high cost or risk associated with false-positive test results.
Mycoplasma-like Organisms (MLOs) INTRODUCTION
MLOs were first observed by electron microscopy by Doi et al. (1967). Despite many attempts, MLOs have not been cultured so far. An inability to culture MLOs made the preparation of specific reagents difficult. As a result, microscopy was the only method for diagnosis of MLO infections. Using partially purified MLO preparations, polyclonal antibodies were produced against some MLOs, however, the antiserum had to be extensively cross-absorbed with antigen preparations from healthy plant tissues to retain some specificity versus MLOs. From 1985 on, the application of hybridoma technology to the production of MLO-specific monoclonal antibodies has greatly benefited both detection and characterization of this group of mollicutes by overcoming the problems encountered with polyclonal antisera. Indeed, for the first time, highly specific reagents were available for MLOs. The first MAbs were produced against a New Jersey strain of aster yellow MLO, using partially purified salivary gland preparations from infected leafhopper vectors as the immunogen (Lin and Chen, 1985). Because the insect vectors of many MLOs are unknown, a more general procedure was developed in which infected plant extracts were the immunogen (Martin-Gros et al, 1987). Since then, MAbs against a large number of MLOs have been obtained and serological techniques similar to those described next have been applied to the diagnosis of MLO infections and to the identification of insect vectors (Fos et al, 1992). DAS-EUSA: Double Antibody Sandwich Enzyme-Linked Immunosorbent Assay MATERIALS Equipment
Same equipment as for C-ELISA with the addition of the following: Incubator, 37°C
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Refrigerator, 4°C Mortars and pestles or Polytron homogenizer Glass homogenizer (for insects) Cheesecloth Buffer
Coating buffer, PBS buffer, and wash buffer; see C-ELISA Sample buffer: the PBS buffer can be supplemented with Tween 20 (see PBSTween) and 2% (w/v) polyvinylpyrrolidone (PVP) MW 25,OCX) Conjugate buffer: PBS-Tween-PVP-ovalbumin: PBS-Tween-PVP containing 0.2% ovalbumin. Substrate buffer: Diethanoloamine, 98 ml; H2O, 800 ml; NaN3, 0.2 g; adjust pH to 9.8 with HCl. Make up to 1 liter with H2O. Reagents
Purified MAb (capture MAb): diluted in coating buffer (the optimal final concentration has to be determined. It ranges generally from 1 to 20 |xg per ml according to the MAb). Alkaline phosphatase(AP)-labeled MAb: diluted in conjugate buffer (the optimal dilution has to be determined. It ranges from 1/500 to 1/4000). Enzyme substrate: P-Nitrophenyl phosphate, 1.0 mg per ml of substrate buffer. PROCEDURES
1. Purification of MAbs and their labeling with alkaline phosphatase are done according to standard protocols. 2. Preparation of the plant extracts: 1 g of leaf midribs is ground in 2 ml of sample buffer in a mortar or with a Polytron homogenizer. The homogenate is filtered through four layers of cheesecloth; 150 jxl are used directly for ELISA. Dilutions of the homogenate can also be made. 3. Preparation of insect extracts: single insects or batches of 5 to 10 insects (leafhopper or psyllids) according to their size and or availability are gound in 300 |xl of PBS in a glass homogenizer. After decantation, the homogenate is pipetted directly in the ELISA plate. DAS-ELISA
1. Add 150 |xl of purified IgG in coating buffer to each well of the microtiter plate. Incubate for 2 to 4 hours at 37°C. Empty plate by shaking. 2. Wash by flooding wells with PBS-Tween, three times for 5 minutes. Empty plate by shaking.
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3. Add 150 |xl of sample extract into duplicate wells. Incubate overnight at 4°C. 4. Wash plate three times with PBS-Tween as in step 2. 5. Add 150 |xl of AP-labeled IgG in conjugate buffer to each well. Incubate for 2 to 4 hours at 37°C. 6. Wash plate three times with PBS-Tween as in step 2. 7. Add 150 |xl of freshly prepared substrate in substrate buffer to each well. Incubate at room temperature for 30 to 60 minutes. 8. Results can be assessed by visual observation of the color (yellow color for positive samples, no color for negative samples) or quantitatively determined by measurement of the absorbance at 405 nm. 9. Controls to be used: Negative controls consist in the replacement of the sample extract by sample buffer alone, with all other steps being the same. Extracts from healthy plants and/or insects of the same species should also be included in the assay. The optical density of the negative control wells should be under 0.1. Positive controls consist of extracts from plants or insects known to be infected with the appropriate MLO. The optical density of such wells is generally high (>2.0).
Indirect Immunofluorescence on Plant Tissue Sections MATERIALS Equipment
Razor blades Freezing microtome Glass slides and coverslips (slides with 10 wells are recommended, CML laboratories) Tweezers Pasteur pipettes Hot plate (55°-60°C) Binocular dissecting microscope Epifluorescence microscope equipped with filters for fluorescein isothiocyanate (FITC) (Ziess III RS with the filter combination BP 455-490/Fr 510/LP 420, for example). Reagents
Mounting solution: glycerol, 50% in PBS FITC-labeled goat anti-mouse immunoglobulin 1% Evans blue.
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PROCEDURES
1. Cut leaf midribs or petioles in 2- to 5-mm-long fragments with a razor blade. 2. Cut longitudinal sections (10 to 30 |xm thick) with a freezing microtome. Transverse sections can also be made but if the number of MLOs is low, they will be more easily detected in longitudinal sections. 3. Spread the sections on a glass slide with the help of tweezers under a binocular dissecting microscope. Remove excess water with a Pasteur pipette or by blotting with paper. 4. Fix the sections by putting the slide on a hot plate at 60°C for 1 hour. Other fixation methods can be used (acetone, methanol) according to the MLO and MAb considered. 5. Incubate the sections with undiluted hybridoma supernatant for 30 minutes at room temperature. 6. Wash three times with PBS containing 0.05% Tween 20. 7. Incubate the sections for 30 minutes in the dark with FITC-labeled goat anti-mouse IgG diluted 100-fold in PBS containing a 100-fold dilution of 1% Evans blue. 8. Wash as in step 6. 9. Mount the sections in the mounting solution. 10. Observe the sections under the epifluorescence microscope. Controls: Negative controls are sections from healthy plants treated in the same way and sections from healthy and infected plants in which step 5 has been modified by replacing hybridoma supematants with PBS, preimmune, or unrelated serum. No specific fluorescence should be observed. Autofluorescence will be seen. Positive controls are sections from MLO-infected plants treated in the same way. DISCUSSION
The DAS-ELISA developed by Clark and Adams (1977) for detection of plant viruses is the most widely used method for MLO detection in both plants and insects. It requires the purification of immunoglobulins (Ig) from hybridoma supematants or ascites fluids and their labeling with an enzyme (generally alkaline phosphatase). When this is done, it allows the processing of a large number of samples very quickly; however, Fos et al. (1992) demonstrated that some plant extracts could inhibit that assay which thus failed to detect the MLO. In this case, indirect immunofluorescence on sections has to be used. This test is very simple and does not require any Ig purification. In addition, IgM can be used much more efficiently than ELISA and results are obtained within a few hours. Moreover, the restricted location of MLOs to phloem tissues makes their detec-
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tion very easy and specific. A freezing microtome and an epifluorescence microscope are necessary for this technique which is, however, sometimes difficult to apply to insects because of the high level of autofluorescence associated with some insect tissues. Other serological methods such as the dot-blot immunobinding assay (Boudon-Padieu et al., (1989) or immunosorbent electron microscopy have also been used for MLO detection; they are not described here as they have not been used extensively.
References Ball, H. J., Mackie, D. P., Finlay, D., McNair, J., and Pollock, D. A. (1990). An antigen capture ELISA test using monoclonal antibodyies for the detection of Mycoplasma californicum in milk. Vet. Immunol. Immunopathol. 25, 269-278. Boothby, J. T., Mueller, R., Jasper, D. E., and Thomas, D. E. (1986a). Detecting Mycoplasma hovis in milk by enzyme-linked immunosorbent assay, using monoclonal antibodies. Am. J. Vet. Res. 47, 1082-1084. Boothby, J. T., Jasper, D. E., and Thomas, C. B. (1986b). Experimental intramammary inoculation with Mycoplasma bovis in vaccinated and unvaccinated cows: Effect on the mycoplasmal infection and cellular inflammatory response. Cornell Vet. 76, 188-197. Boudon-Padieu, E., Larrue, J., and Caudwell, A. (1989). ELISA and dot blot detection of flavescence doree-MLO in individual leafhopper vectors during latency and inoculative state. Curr. Microbiol. 19, 357-364. Clark, M. P., and Adams, A. N. (1977). Characteristics of the microplate method of enzyme-linked immunosorbent assay for the detection of plant viruses. J. Gen. Virol. 34, 475-483. Doi, Y., Teranaka, M., Yora, K., and Asuyama, H. (1967). Mycoplasma or PLT group like microorganism found in the phloem elements of plants infected with mulberry dwarf, potato witches'broom, aster yellows or paulownia witches'broom. Ann. Phytopathol. Soc. Jpn. 33, 259-266. Feldman, R. C , Henrich, B., Kolb-Bachofen, V., and Hadding, U. (1992). Decreased metabolism and viability oiMycoplasma hominis induced by monoclonal antibody-mediated agglutination. Infect. Immun. 60, 166-174. Fos, A., Danet, J. L., Zreik, L., Gamier, M., and Bove, J. M. (1992). Use of a monoclonal antibody to detect the stolbur mycoplasmalike organism in plants and insects and to identify a vector in France. Plant Dis. 76, 1092-1096. Heller, M., Barthold, E., Pfuntzner, H., Leirer, R., and Sachse, K. (1993). Antigen capture ELISA using monoclonal antibodies for the detection of Mycoplasma bovis in milk. Vet. Microbiol. 37, 127-133. Lin, C. P., and Chen, T. A. (1985). Monoclonal antibodies against the aster yellows agent. Science 227, 1235-1236. Martin-Gros, G., Gamier, M., Iskra, M. L., Gandar, J., and Bove, J. M. (1987). Production of monoclonal antibodies against phloem limited prokaryotes of plants: A general procedure using extracts from infected periwinkles as immunogen. Ann. Inst. Pasteur Microbiol. 138, 625637. Morsy, M. A., Panagala, V. S., Gresham, M. M., andToivio-Kinnucan, M. (1992). Acoagglutination assay with monoclonal antibodies for rapid laboratory identification of Mycoplasma synoviae. Avian Dis. 36, 149-153.
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Sorensen, V., Barfod, K., and Feld, N. C. (1992). Evaluation of a monoclonal blocking ELISA and IHA for antibodies to Mycoplasma hyopneumoniae in SPF-pig herds. Vet. Rec. 130, 488-490. Thomas, C. B., Jasper, D. E., Boothby, J. T., and Dellinger, J. D. (1987). Enzyme-linked immunosorbent assay for detection of Mycoplasma californicum-spQcific antibody in bovine serum: Optimization of assay determinants and control of serologic cross-reactions. Am. J. Vet. Res. 48, 590-595.
B7 MICROIMMUNOFLUORESCENCE David Taylor-Robinson
introduction The immunofluorescence test described in this chapter is concerned with the detection and measurement of antibody and not with the serological identification of mycoplasmas, where the test is usually applied to colonies on agar. The latter has been described in detail by Gardella et al. (1983). A variety of methods is available for detecting mycoplasmal antibodies in sera and other body fluids, but not all are sufficiently sensitive for quantitative measurements (TaylorRobinson, 1989). Fluorescent antibody staining of mycoplasmas sedimented from broth cultures and fixed to glass slides was described in the early 1960s (Clark et al., 1963). Soon after. Mycoplasma hominis sedimented from a broth culture and fixed to glass slides was used as an antigen in an immunofluorescence test to demonstrate a serological response in a patient with a febrile gynecological disease from whose blood this mycoplasma had been recovered (TuUy et al, 1965). After a long fallow period, indirect microimmunofluorescence (MIF) on slides was used to measure antibody to M. pneumoniae (Sillis and Andrews, 1978) and then to M, genitalium (Purr and Taylor-Robinson, 1984); antibody responses to the latter were detected in men with nongonococcal urethritis (Taylor-Robinson et a/., 1985) and in women with salpingitis (M0ller et a/., 1984). Indeed, this indirect MIF test is sufficiently sensitive, specific, and simple to be of value.
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Media and Antigen Preparation The indirect MIF test may be applied to measuring antibodies to any of the members in the class MoUicutes. Hence, the medium used for the production of antigen is that which will provide optimal growth of the mycoplasma under consideration. The basic composition of culture media has been presented elsewhere (Freundt, 1983; Shepard, 1983; Whitcomb, 1983; see also Chapters A2 and A3 in Vol. I). Antigen is prepared usually from organisms grown in liquid medium rather than from mycoplasmal colonies scraped from the surface of agar medium. Medium, after incubation at the appropriate temperature for multiplication of the mycoplasmal organisms in question, is centrifuged. Since the organisms are usually in aggregates, it is often possible to deposit them by centrifugation on a bench centrifuge. The deposit is resuspended in 10-20 ml of phosphate-buffered saline (PBS; 0.01 M, pH 7.2) and recentrifuged; this constitutes the first washing. After two further washings in PBS, the deposit is resuspended in a small volume of PBS. This antigen is then stored in aliquots of 0.1 ml at -20°C or at a lower temperature.
Procedure The indirect MIF test is undertaken as follows: (i) an aliquot of antigen is thawed and diluted in PBS so that it contains 0.5-1.0 mg of protein per ml for use; (ii) the antigen is applied with a mapping pen to a clear area on a Tefloncoated 3 inch X 1 inch glass slide (Thomas et al., 1976); the number of "spots" of antigen to be placed on each slide is optional but it might be reasonable to have two rows of 12 spots along the length of the slide; (iii) the antigen spots are allowed to dry at room temperature and then they are "fixed" in acetone for 30 minutes; (iv) the serum or other body fluid is diluted in serial twofold steps in PBS (for example, dilutions from 1:2 to 1:512) in U-shaped, polystyrene, microtiter plates and then a 15-|xl volume (one drop) of each dilution is added to the antigen spots with a Pasteur or Eppendorf pipette and the slide is incubated at 37°C for 30 minutes in a moist chamber; (v) the slide is washed in PBS three times (10 minutes each), then held vertically and excess fluid is blotted away from the edge, and the slide is allowed to dry at room temperature; (vi) fluorescein-conjugated (FC) anti-human globulin (if human sera are being tested; anti-bovine if bovine sera are being tested, etc.) prepared in sheep or other species is added in a 15-|xl volume to each antigen spot except one; thus, there will be nine spots of antigen to which serum dilutions and an FC antiglobulin have been added, two spots of antigen to which FC antiglobulin has been added, and one spot of antigen alone; and (vii) the slide is incubated at 37°C for 30
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minutes in a moist chamber, washed three times in PBS, dried at room temperature, and examined by epifluorescence microscopy (100 and 400 x magnification) (see Gardella et aL, 1983, for details of microscopic equipment). The titer of the serum or other fluid being tested is the highest dilution at which fluorescence is detected.
Discussion The indirect MIF test described in this chapter is similar to that used for the quantitative measurement of antibody to Chlamydia trachomatis (Thomas et al., 1976). The concentration of protein in the antigen preparation mentioned earlier is not a crucial requirement for a successful test. However, there should be sufficient antigen to enable fluorescence to be detected easily and knowing the protein content is helpful when preparing another batch of antigen so that there is standardization. It is also important to incorporate a standard positive serum and a negative serum as controls in each batch of samples tested. The indirect MIF test has several advantageous features: (i) reproducibility can be maintained because it is possible to store a large batch of antigen in aliquots that are frozen so that there should be no variation from one aliquot to another, (ii) reproducibility can also be maintained by prepreparing multiple slides with antigen spots and storing them at 4° or — 20°C, (iii) only small volumes of reagents are required, (iv) estimation of IgG, IgM, or IgA antibodies is accomplished easily by using the appropriate antiglobulins, (v) antibiotics in serum do not produce spurious results in the same way that they might in a metabolisminhibition test where the inhibitory effect of an antibiotic might be mistaken for that of antibody, and (vi) the test is quick to perform, particularly if slides with antigen spots have been preprepared, and it is relatively easy to read. It has to be recognized, however, that the reading of the test is subjective and a decision has to be made as to whether the end point is the highest dilution of serum at which there is fluorescence equal to the strong fluorescence seen at the lower dilutions or whether fluorescence of lesser brightness is taken as the end point. It is important that whatever decision is taken, it should be maintained so that there is consistency throughout the test and from one test to another. The indirect MIF test has been used to measure antibodies directed against several mycoplasmas, including M. hominis, as mentioned before, M. pneumoniae (Wreghitt and Sillis, 1985; Sillis, 1990), andM. synoviae of avian origin (Bradbury et al, 1990). It has been particularly useful in seeking serum antibodies to M. genitalium where the sensitivity of the procedure was found to be greater than that of the metabolism-inhibition test (Furr and Taylor-Robinson, 1984). The specificity of the indirect MIF test is probably attributable to the fact
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that antibody to surface components of the mycoplasmal membrane is detected. This provides the rationale for using whole organisms as the antigen rather than disintegrating them by sonication in distilled water.
References Bradbury, J. M., McCarthy, J. D., and Metwali, A. Z. (1990). Microimmunofluorescence for the serological diagnosis of avian mycoplasma infections. Avian Pathol. 19, 213-222. Clark, H. W., Bailey, J. S., Fowler, R. C , and Brown, T. M. (1963). Identification of Mycoplasmataceae by the fluorescent antibody method. J. Bacteriol. 85, 111-118. Freundt, E. A. (1983). Culture media for classic mycoplasmas. In "Methods in Mycoplasmology" (S. Razin and J.G. TuUy, eds.). Vol. 1, pp. 127-135. Academic Press, New York. Furr, P. M., and Taylor-Robinson, D. (1984). Microimmunofluorescence technique for detection of antibody io Mycoplasma genitalium. J. Clin. Pathol. 37, 1072-1074. Gardella, R. S., Del Giudice, R. A., and TuUy, J. G. (1983). Immunofluorescence. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.), Vol. 1, pp. 431-439. Academic Press, New York. M0ller, B. R., Taylor-Robinson, D., and Furr, P. M. (1984). Serological evidence implicating Mycoplasma genitalium in pelvic inflammatory disease. Lancet i, 1102-1103. Shepard, M. C. (1983). Culture media for ureaplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.). Vol. 1, pp. 137-146. Academic Press, New York. Sillis, M. (1990). The limitation of IgM assays in the serological diagnosis of Mycoplasma pneumoniae infections. J. Med. Microbiol. 33, 253-258. Sillis, M., and Andrews, B. (1978). A simple test forM. pneumoniae IgM. Zentralbl. Bakteriol., Parasitenkd., Infektionskr. Hyg., Abt. 1: Orig., Reihe A 241, 239-240. Taylor-Robinson, D. (1989). Genital mycoplasma infections. Clin. Lab. Med. 9, 501-523. Taylor-Robinson, D., Furr, P. M., and Hanna, N. F. (1985). Microbiological and serological study of non-gonococcal urethritis with special reference to Mycoplasma genitalium. Genitourin. Med. 61, 319-324. Thomas, B. J., Reeve, P., and Oriel, J. D. (1976). Simplified serological test for antibodies to Chlamydia trachomatis. J. Clin. Microbiol. 4, 6-10. TuUy, J. G., Brown, M. S., Sheagren, J. N., Young, V. N., and Wolff, S. M. (1965). Septicemia due to Mycoplasma hominis type 1. N. Engl. J. Med. 273, 648-650. Whitcomb, R. F. (1983). Culture media for spiroplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.). Vol. 1, pp. 147-158. Academic Press, New York. Wreghitt, T. G., and SUUs, M. (1985). A |x-capture ELISA for detecting Mycoplasma pneumoniae IgM: Comparison with indirect immunofluorescence and indirect ELISA. J. Hyg. 94, 217227.
B8 IMMUNOBLOTS AND IMMUNOBINDING David Thirkell and Bernard L. Precious
A number of techniques utilize antibodies for the detection of or to gain information on antigens. Many require a means of visualization and some require ancillary techniques [e.g., radiolabeling of mycoplasma cells; sodium dodecyl sulfate-Polyacrylamide Gel Electrophoresis (SDS-PAGE)]. This chapter is not comprehensive, it deals only with protein antigens and it introduces the reader to the principles by using selected examples and gives typical protocols. For most, more comprehensive details may be gained from one of the many laboratory manuals available (e.g., Coligan et al., 1994).
Immunoblotting (Western Blotting/Dot-Blotting) Introduction Immunoblotting involves the immobilization of antigens on a membrane of nitrocellulose, nylon, or polyvinylidine difluoride (PVDF). Antigens are then detected using specific polyclonal antiserum (e.g., 50 |xl diluted to 20 ml in mycoplasma growth medium to prevent detection of medium components), monospecific or monoclonal antisera, and a means of visualization with either enzymatic or radioactive techniques. In a simple system (dot-blotting), antigens may simply be "dropped" (5 |xl applied directly) onto the nitrocellulose and allowed to absorb. However, complex antigen mixtures are often first separated by SDS-PAGE and transferred to, e.g., nitrocellulose by Western blotting be151 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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fore visualization of data. This technique has the advantage that molecular weights of antigens may be determined. A typical protocol is outlined next. Further information may be gained on specific antigens using both isoelectric focusing and two-dimensional PAGE in conjunction with blotting.
Materials
Mycoplasma cells Denature mix [e.g., 5 M urea, 3.5 M 2-mercaptoethanol, 2% (w/v) sodium dodecyl sulfate (SDS), 0.1% (w/v) bromophenol blue] Boiling water bath/Eppendorf tubes SDS-PAGE equipment complete with appropriate power pack Preprepared SDS-PAGE chemicals and polymerizing reagents (commercially available) Prestained protein molecular weight markers (e.g., Sigma) Nitrocellulose sheet (cut to the size of the gel used for SDS-PAGE) Semidry blotting equipment (e.g., Bio-Rad, Pharmacia, Ancos) and associated power pack; anode and cathode buffers; dialysis membrane, if required (see manufacturer's instructions) Whatman No. 1 filter paper (cut to appropriate size) Flat-topped shaker Phosphate-buffered saline (PBS); PBS containing 0.05% (v/v) Nonidet P-40 (PBS-N) Tris-buffered saline (TBS) and TBS-0.05% (v/v) Nonidet P-40 (TBS-N) 3% (w/v) bovine serum albumin (BSA) in PBS Antibody (polyclonal antiserum, monospecific or monoclonal antiserum) A method for visualizing the result of the immunoblot (see Procedures)
Procedures SDS-PAGE (MOUCHES AND BOVE, 1983; SEE ALSO CHAPTER D4, VOL. I)
The nature of the separating gel used is dictated by the molecular weight range of the antigens to be electrophoresed. Mycoplasma samples to be electrophoresed (e.g., 25 |jLg mycoplasma protein in 50 |xl per lane on a 20 x 20-cm gel) and molecular weight markers are boiled individually (Eppendorf tubes; 3 minutes) after the addition of an equal volume of denature mix. After cooling, samples and the molecular weight markers are loaded onto separate gel lanes (e.g., with a Hamilton syringe). The gel is electrophoresed at 50 V (constant voltage) overnight or for shorter periods at 200 V if minigels are being electro-
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phoresed. Bromophenol blue in the sample enables the "front" to be detected and this should not be allowed to run off the bottom of the gel. ELECTROPHORETIC TRANSFER OF PROTEINS/POLYPEPTIDES TO INERT SUPPORT
The most commonly used technique is semidry blotting, although wet and passive blotting are alternatives. Semidry blotting is quick and gives good resolution, wet blotting takes longer but may give better renaturation of components, and passive blotting is the longest to perform, gives the greatest renaturation, but has the poorest resolution. As an example, a typical protocol for semidry blotting is described. Semidry blotters have two flat electrodes; an anode (lower plate) and a cathode (upper plate). Essentially, a stack is made on the anode plate consisting of several sheets of filter paper (soaked in anode buffer/buffers), a sheet of nitrocellulose (presoaked in distilled water), the gel (after SDS-PAGE) and then six sheets of filter paper (soaked in cathode buffer). All air must be expelled between the individual layers of the stack by gently rolling a pipette over the wet surface or uneven transfer of electrophoresed material will occur. The cathode plate is placed on top of the final stack and the electrophoretic transfer to the nitrocellulose sheet is performed with a constant current of 0.8 mA per cm^ of gel (20°C, 1 hour). A protocol is normally included by the manufacturer for the simultaneous electrophoretic transfer of polypeptides from more than one gel to separate sheets of nitrocellulose (dialysis membranes required). DETECTION OF ANTIGENS
The lane containing the prestained molecular weight markers demonstrates transfer efficiency to the nitrocellulose and is used to determine the apparent molecular weights of any antigens detected. The nitrocellulose sheet is removed from the semidry blotter, washed well with PBS, and the unimpregnated surface area is "blocked" with 3% BSA in PBS on a shaker (20°C, 1 hour). After thorough washing of the nitrocellulose sheet with PBS-N, the sheet as a whole, or after cutting into appropriate strips, is incubated (e.g., 37°C, 1 hour) with antibody [e.g., homologous/heterologous polyclonal antiserum (e.g., diluted 1:500 (v/v) in mycoplasma growth medium to prevent detection of medium components), monospecific antiserum, or MAb at a predetermined dilution]. After thorough washing with PBS-N, antigens are recognized by one of several methods. Use of ^^^l-Labeled Protein A (Specific Activity 30 mCi mg~^)
The nitrocellulose sheet/strips are agitated on a flat-topped shaker (20°C, 1 hour) in 15 ml PBS-N to which ^^sj.iabeled protein A (around 5 fxCi per lane
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on the original gel) is added. The nitrocellulose is washed thoroughly with PBSN, "dabbed" dry, and exposed to X-ray film (with an intensifying screen), usually overnight at --70°C. Antigenic bands are readily seen after development of the X-ray film. Use of Enzyme-Linked Detection Systems
A second antibody (anti-IgG or anti-Ig), which must have been raised against antibodies of the animal species used to raise the first antibody/antibodies (e.g., anti-mouse/anti-rabbit), is linked to either alkaline phosphatase or, more commonly, to horseradish peroxidase. Alternatively, either one of these enzymes can be linked to protein A. For a horseradish peroxidase-linked second antibody, a typical procedure is as follows. Sheets or strips of nitrocellulose are incubated (20°C, 3 hr) with an appropriate dilution of second antibody [in TBS containing 3% (w/v) BSA]. After washing with TBS (20°C, 10 minutes), 2x with TBS-N (20°C, 10 minutes), and then with TBS alone (20°C, 10 minutes), the nitrocellulose is developed in the dark (up to 1 hour) in 60 mg 4-chloro-l-naphthol in 20 ml methanol and 80 ml TBS to which 60 |JL1 30% (v/v) H2O2 is added immediately before use. Antigens are seen as purple bands. A typical immunoblot of strains of Mycoplasma ovipneumoniae using a polyclonal antiserum raised in rabbit against one of the strains is seen in Fig. 1.
O N O N O V O O V X - J — J — J — I c D
> o r - - C L Q _ a . Q _ a . Q L a . Q L V D
-18-/. Fig. 1. Immunoblot of 22 strains of Mycoplasma ovipneumoniae probed with a polyclonal antiserum raised in rabbit against strain 956/2 and a horseradish peroxidase-linked anti-rabbit IgG (Reproduced from Thirkell et al., 1990a, Vet. Microbiol. 21, 241-254, by permission of Elsevier Science Publishers.
B8 Immunoblots and Immunobinding Enhanced Chemiluminescence (ECL, Amersham) Method of Visualization Instructions Supplied by the Manufacturer)
155 (Full
A horseradish peroxidase-linked second antibody is used as described earlier (an alternative is the use of peroxidase-linked protein A), but development is on X-ray film and is dependent on chemiluminescence. It is a rapid technique which gives very good resolution of antigenic material.
Discussion Despite the variations in protocol, the method used for visualization of the final result(s) is important. With ^^^I-labeled protein A, only immunoglobulins G are detected. It is possible to overcome this by adding an additional step. The blot may be incubated with a second antibody (of anti-IgG but not enzymelinked) before application of the protein A. If prolonged exposure of the autoradiogram at — 70°C is required, the bands detected can be diffuse. Using the horseradish peroxidase-linked second antibody, tight bands are seen on a clean background but the sensitivity of the visualization procedure is not good. The new ECL visualization is the method of choice. Tight bands are seen on the X-ray film and the sensitivity is usually close to that achieved with ^^sj.iabeled protein A.
Immunoprecipitation Introduction Immunoprecipitation is used for detecting nondenatured epitopes. It often (but not necessarily) depends on the interaction of radiolabeled solubilized antigens and antibodies (IgG) to form a complex which is recovered following binding to immobilized protein A. The radiolabeled antigen is then detected by autoradiography following SDS-PAGE of the complex. [^^S]Methionine is often the metabolic label of choice. However, some mycoplasmas do not incorporate this isotope well. Surface components of such mycoplasmas may be labeled with 1251; cytosolic components can also be labeled after lysis of the cells. It should also be noted that this radiolabeling technique will also label any contaminating medium components and should thus be used with care. If an unlabeled antigen is used, an alternative visualization procedure is employed (see Immunoblotting).
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Materials
[^^SJMethionine or i^^I-labeled Bolton and Hunter reagent (A^-succinimidyl 3-(4hydroxy-5-[i25j] iodophenyl)propionate) (specific activity 2000 mCimmolO 0.1 M borate buffer, pH 8.3 0.1 M glycine in 0.1 M borate buffer, pH 8.3 Antiserum or monoclonal antibody (MAb) 10% (w/v) fixed suspension of Cowan strain A Staphylococcus aureus (Kessler, 1975) or 100 mg ml-i protein A-Sepharose CL-4B in PBS Phosphate-buffered saline Sucrose Lysis/immunoprecipitation buffer (IPB): 10 mM Tris-HCl, pH 7.2, 5 mM EDTA, 0.5% (v/v) Nonidet P-40, 0.65 M NaCl, 0.1% (w/v) NaN3, 1 mM phenylmethylsulfonyl fluoride (added just before use) Centrifuge SDS-PAGE equipment and chemicals Dimethyl sulfoxide (DMSO) Gel soak: 22% (w/v) 2,5-diphenyloxazole (PPO) in DMSO (Laskey and Mills, 1975) or Amplify (Amersham) Vacuum gel drier X-ray film
Procedures RADIOLABELING OF MYCOPLASMA CELLS Metabolic Labeling with P^SjMethionine (Preferred Method)
Generally, mycoplasma cultures (usually 100 ml) are grown to a midlogarithmic phase and cells are pelleted (20,000 g, 4°C, 5 minutes), then suspended in one-tenth original volume in a methionine-depleted or methionine-free tissue culture medium to which 300 |xCi[^^S]methionine is added. After incubation (37°C, 2 hours), cells are pelleted as described earlier and may be subjected to a 2- to 3-hour "cold chase" (37°C) by replacement of tissue culture medium with normal mycoplasma medium; after centrifugation, they are washed (3x) with PBS and the final pellet is suspended in IPB. Alternative Surface Labeling of Mycoplasma Cells with '^^/ (Precious et ai, 1987)
Washed pellets from fresh mycoplasma cultures are suspended in 0.1 M borate buffer (pH 8.3), centrifuged as described earlier, and resuspended in the same buffer at approximately 10^ CCU (color changing units) ml-^ Cells (0.1 ml) are
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radiolabeled (4°C, 15 minutes) with 100 |xCi i^sj.iabeled Bolton and Hunter reagent according to the supplier's instructions (Amersham). The reaction is terminated by the addition of 0.1 M glycine in 0.1 M borate buffer (pH 8.3). The cells are washed (3x) in PBS to remove any labeled internal components released by autolysis, resuspended in IPB, and stored at -20°C. Labeling of Cytosolic
Components
Mycoplasma cells prepared as described earlier are first lysed with IPB and are then labeled as just described. IMMUNOPRECIPITATION WITH LABELED MYCOPLASMA CELLS
Radiolabeled mycoplasma cells in IPB are sonicated on ice (4 x 10 seconds with cooling), and after centrifugation (30,000 g, 4°C, 10 minutes), the supernatant is collected. Five microliters of undiluted polyclonal antiserum or 5 |JL1 of undiluted ascitic fluid is added to 50 |xl of supernatant and the mixture is incubated (4°C, 1 hour). A fixed suspension (100 jxl) of Cowan strain A Staphylococcus aureus or 75 JJLI protein A-Sepharose CL-4B is added, and after incubation (4°C, 1 hour) and centrifugation (600 g, 4°C, 10 minutes), the pellet is resuspended in a small volume of IPB in 10% (w/v) sucrose. The pellet is washed (3x) in this buffer and sucrose mixture, and the final pellet is suspended in 100 |JL1 of the "denature mix" (see Immunoblotting) and boiled (3 minutes). After centrifugation (600 g, 4°C, 10 minutes), the supernatant is subjected to SDSPAGE. After electrophoresis, the gel is fixed in "gel fix" (20 minutes). Then, if the 1251 label is used, the gel is vacuum dried (60°-80°C). If the 35S label is used, the gel is washed (3 x 30 minutes) in 50 ml DMSO, soaked in 22% (w/v) PPO in DMSO (gel soak; 3 hours) (safety aspects must be considered), and then washed with running water (30 minutes) and vacuum dried (60°C). The dried gel is subjected to autoradiography or fluorography [X-ray film, -70°C, overnight to 48 hours in a cassette with an intensifying screen (i^sj-iabeled cells only)]. A typical result is seen in Fig. 2 using [^^SJmethionine-labeled Mycoplasma ovipneumoniae, a homologous polyclonal antiserum, and two monoclonal antibodies. Discussion
1251 surface labeling can also be used to identify surface antigenic components, e.g., the 96-kDa antigen of Ureaplasma urealyticum (Precious et ai, 1987). Problems with i25i.iabeled contaminants can be minimized by using higher volumes of antisera diluted in growth medium to "swamp out" antibodies against medium components or by substituting fetal calf serum for horse serum in the mycoplasma growth medium.
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David Thirkell and Bernard L. Precious
—
-WkDa
Fig. 2. Immunoprecipitation of [^ssjmethionine-labeled M. ovipneumoniae (strain 956/2) with homologous polyclonal serum (B) and with two MAbs raised against the organism (C/D). (A) An autoradiogram of the radiolabeled cells alone (Reproduced from Thirkell et a/., 1990a, Vet. Microbiol. 21, 241-254, by permission of Elsevier Science Publishers.
The technique may also be used for the detection of acylated surface-expressed antigens after growth of the mycoplasma in the presence of ^H-labeled palmitic acid (see Chapters C2 and C3, Vol. I). Autoradiograms of such results normally require prolonged exposure at -70°C. Immunoaffinity Chromatography Introduction
Enriched MAbs recognizing a specific antigen may be coupled to an inert support and an immunoaffinity column prepared with the complex. Such columns allow a rapid single-step purification of antigens.
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Materials Enriched MAb (as detailed later) Cyanogen bromide-activated Sepharose 4B IMHCl Centrifuge PBS (see Immunoblotting) and PBS with 0.2% (v/v) Nonidet P-40 (PBS-N') Naphthalene black stain [0.2% (w/v) naphthalene black in 10% (v/v) glacial acetic acid, 40% (v/v) methanol, 50% (v/v) distilled water] 1- and 2-ml columns "End-over-end" shaker Coupling buffer: 0.1 M NaHCOa, pH 8.3, containing 0.5 M NaCl "Blocking" buffer: 0.2 M NaHCOs, 0.1 M NaCl, pH 8.0, containing 1 M ethanolamine Sintered glass funnel 0.1 M acetate buffer, pH 4.0 Elution buffers, e.g., 0.1 M glycine-HCl, pH 2.8, or 0.1 M borate buffer, pH 10.0 1 M Tris-HCl, pH 7.5 Dialysis tubing
Procedures ENRICHMENT OF MONOCLONAL ANTIBODY/ANTIBODIES Immunoglobulin Class G (IgG)
Protein A-Sepharose is swollen overnight in PBS at 4°C (300-mg beads ml-i buffer) after which a minicolumn is made in, e.g., a 1-ml disposable syringe. After column equilibration with PBS, 1 ml of ascitic fluid is loaded and the column is washed with PBS-N' until no further protein is eluted (assayed by dotting 5-|xl aliquots of column eluate onto a nitrocellulose sheet and staining with naphthalene black). The immunoglobulin is eluted with 0.1 M glycine-HCl (pH 2.8), and 400-|xl fractions are collected in Eppendorf tubes (containing 100 |JL1 1 M Tris-HCl, pH 7.5). Elution is continued until no more protein is released (assayed as described earlier; usually a maximum of 24 fractions are required). High protein-containing fractions are pooled and dialyzed overnight against PBS. The Ig class may be proved by immuno-dot-blot using a horseradish peroxidase-linked anti-IgG antibody (see Immunoblotting). A few drops of 0.1% (w/v) sodium azide are added at this stage if the enriched MAb is to be kept.
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David Thirkell and Bernard L. Precious
MAb Not of Immunoglobulin Class G
One milliliter of ascitic fluid is precipitated by dropwise addition (with stirring) of saturated (NH4)2S04 to a final concentration of 50% (w/v). After centrifugation (5000 g, 5 minutes), the pellet is resuspended in PBS and is dialyzed against two changes of PBS overnight at 4°C; this solution is then passed through a Sephadex G-200 column (preswoUen in PBS and packed into a 30 x 1.5-cm column in PBS). The column is washed with PBS and 1.5- to 2.0-ml fractions are collected (checked for protein as before). Although both procedures produce enriched MAb in PBS, protein A Sepharose gives a more pure preparation. Before use, MAbs are dialyzed overnight at 4°C against coupling buffer. PREPARATION OF IMMUNOAFFINITY COLUMN
One gram of cyanogen bromide-activated Sepharose 4B is swollen in 200 ml 1 mM HCl and is then filtered onto a sintered glass funnel where it is washed with 250 ml deionized water followed by 5-10 ml coupling buffer. The washed Sepharose is removed into 10 ml coupling buffer and 1 ml enriched MAb is added. The mix is placed into a small sealed container and is shaken overnight at 4°C using an end-over-end shaker. After centrifugation (1000-2000 g, 5 minutes), the remaining reactive sites on the Sepharose are "blocked" by suspension in 20 ml blocking buffer and incubation (20°C, 2 hours). The complex is filtered onto the sintered glass funnel, and excess uncoupled MAb and blocking buffer are removed by alternate washing with 0.1 M acetate buffer (pH 4.0) and coupling buffer (pH 8.3), using 50-60 ml of each buffer. The affinity column complex may be stored in PBS containing 0.1% (w/v) sodium azide; otherwise, a minicolumn is packed in a 2-ml syringe. The minicolumn must be equilibrated before use with the same buffer as used for solubilization of the antigen to be purified. ANTIGEN PREPARATION
The antigen to be purified must be in solution in an appropriate buffer but the method of solubilization will be dependent on the nature of the antigen. It should be kept in mind that agents such as urea or detergents may also disrupt the antibody on the column. PURIFICATION
The antigen solution is loaded onto the column which is washed with loading buffer until no more protein is eluted (test for protein as before). Bound antigen is eluted from the column [e.g., with 0.1 M glycine-HCl (pH 2.8) or with 0.1 M
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borate buffer (pH 10.0), although other antigen/antibody complexes may require more stringent conditions for elution] and 0.4-ml fractions are collected in Eppendorf tubes each containing 100 |JL1 1 M Tris-HCl (pH 7.5). The fractions are assayed for protein (as before) and the major protein-containing fractions are pooled and then dialyzed overnight against PBS. The purified antigen is either used directly or aliquoted and stored at — 20°C.
Discussion
Some antigens are only eluted at high pH, others only at low pH, and others may require more stringent conditions, resulting in denaturation of both antigen and antibody. Consideration of the elution conditions is particularly important if the antigen being purified is an enzyme, as the purified enzyme must be in an "active" form. With highly hydrophobic proteins, it is usually necessary to keep a solubilizing detergent in the eluting buffer to prevent precipitation of the antigen in the column.
Determination of Multiple Epitopes on an Antigen Introduction
Antigenic molecules may express more than one epitope. When a number of MAbs against a single antigen are available, this determination may be undertaken using a competition binding assay.
Materials
MAbs (some enriched from ascitic fluids) 0.02% (w/v) chloramine-T in PBS Nai25i (100 mCiml-i) 0.03% (w/v) sodium metabisulfite "Coarse" Sephadex G-50 Nitrocellulose sheet X-ray film Phosphate-buffered saline with and without 0.2% (v/v) Nonidet P-40 (PBS-N') 84-well Terasaki plates (bases removed; Sterilin) 3% (w/v) BSA in PBS
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David Thirkelt and Bernard L. Precious Terminated reaction + 100)xl PBS
Coarse Sephadex G50 Centrifuge Tube
Eppendorf Tube
Fig. 3.
Syringe column assembly used at the final stage of radiolabeling of enriched MAb.
Procedures RADIOLABELING OF ENRICHED MAb (SEE IMMUNOAFFINITY CHROMATOGRAPHY)
To 20 fxl 0.02% (w/v) chloramine-T in PBS and 500 |xCi Na^^si, 20 yA enriched MAb in PBS (around 5 mg protein m l ^ is added and the mixture is incubated (20°C, 2 minutes). The reaction is terminated by the addition of 20 |xl 0.03% (w/v) sodium metabisulfite. Excess label is removed by centrifugation (300 g, 3 minutes) of labeled MAb through a 1-ml syringe column of coarse Sephadex G-50 (see Fig. 3). TITRATION OF LABELED ANTIBODY
Four-or fivefold dilutions of radiolabeled MAbs are made in a Terasaki plate | L1 wellO- A nitrocellulose sheet (dimensions of the Terasaki plate) is coated (10 U with a sonicated extract of the mycoplasma against which the MAbs have been raised by incubation (20°C, 1 hour). The nitrocellulose is then washed with several changes of PBS, and any uncoated nitrocellulose is blocked with 3% (w/v) BSA in PBS (20°C, 20 minutes). This coated and blocked sheet is then placed on top of the Terasaki plate. An inverted empty Terasaki plate is placed above the nitrocellulose sheet and the "sandwich" is clipped firmly before inversion and incubation (20°C, 1 hour). The "sandwich" is again inverted, and the nitrocellulose sheet is quickly removed and washed extensively (several changes of PBS). The antibody titration end point is determined using one of the visualization methods described (see Immunoblotting). Labeled MAb is used in the competition assay (see later) at a concentration equal to 8-10 x the titration end point [e.g., 1/3200 dilution of MAb in 3% (w/v) BSA in PBS].
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163
COMPETITION ASSAYS
Another coated and blocked nitrocellulose sheet (as just described) is prepared, then washed well with several changes of PBS. Unlabeled MAbs are tested for their ability to compete with the binding of the ^^sj.jabeled MAb by making fourfold dilutions of ascitic fluids (starting at a 1/20 dilution) in the appropriate dilution (see titer earlier) of labeled MAb in PBS containing 1% (w/v) BSA (total volume per well, 15 |xl; Terasaki plate). The coated and blocked nitrocellulose sheet is placed on top of the Terasaki plate and a "sandwich" is made (as earlier). The "sandwich" is clipped together and, after inversion, is incubated (20°C, 1 hour). After removal, the nitrocellulose is washed extensively with PBS-N', dried, and exposed to X-ray film overnight at — 70°C before development of the film (Randall et al., 1987). INTERPRETATION OF ASSAY
Results of two competition assays with anii-Ureaplasma urealyticum urease MAbs are seen in Fig. 4. Lanes A show competition for the binding of labeled MAb with dilutions of the homologous unlabeled MAb. With the i^^I-labeled MAb UU8/29, competition is seen with MAbs B and E which thus recognize the
^20
^20
125
125 I - LABELLED UU8/29
I-LABELLED UU8/5
Fig. 4. Competition assays with ^25|_|abeled Ureaplasma urealyticum anti-96-kDa MAbs and other unlabeled anti-96-kDa MAbs. Fourfold dilutions of unlabeled antibody were made in i25|-|abeled MAbs (UU8/5, UU8/29) (Reproduced from Thirkell etaL, 1990b, by permission of Gustav Fischer Verlag.
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David Thirkell and Bernard L. Precious
same epitope. Groupings of MAbs which recognize a similar epitope are made, and the procedure is repeated with different ^^sj.iabeled MAbs. With UU8/5, competition is only seen with homologous unlabeled MAb and MAb B. Discussion
Through the performance of a series of such competition assays, the minimum number of epitopes on an antigen can be determined. The number of different epitopes found will obviously depend on the number of different MAbs available against the antigen. Results as seen with ^^^lAabdcd UU8/29 in lanes F, G, and H are difficult to interpret. They may be the consequence of true competition between MAbs recognizing the same epitope or they may be the consequence of steric hindrance in the binding of MAbs to two different but adjacent epitopes.
Immunocytochemistry: Immunogold Labeling Introduction
Several techniques are available for the localization of mycoplasma antigens [e.g., labeling with ^^sj for surface-expressed proteins, use of phase partition with Triton X-114 (Bordier, 1981) for hydrophobic (membrane) and hydrophilic (cytosolic) proteins, or preparation of "membranes"]. A more definitive tool is the combination of electron microscopy and immunogold labeling of thin sections of cells (see also Chapter A7 in Vol. I). The mycoplasmas are grown with fetal calf serum instead of with horse serum in the growth medium to minimize cell pellet contamination with serum components. Enriched MAb (IgG; see Immunoaffinity Chromatography) is bound to its antigen before being complexed with protein A-gold (electron-opaque). Fixation of mycoplasmas is required, and since glutaraldehyde cross-links some proteins (especially those surface expressed), thus destroying some epitopes, the basic technique used has to be modified frequently according to the mycoplasma and to the type and location of the antigen being investigated. Three variations have been successful with ureaplasma antigens (Myles et al., 1991) and examples of what is required, are detailed next. Materials
Mycoplasma cells from culture 1% (v/v) and 2.5% (v/v) glutaraldehyde in PBS
B8 Immunoblots and Immunobinding
1 65
Centrifuge 30% (v/v), 50% (v/v), and 70% (v/v) ethanol L.R. White resin (water-soluble resin; medium; Taab); Beem capsules; vacuum oven Electron microscope; ultramicrotome; copper grids (300 nm; Emscope) Buffer A: 20 mM Tris-HCl (pH 8.2) containing 20 mM NaN3, 225 mM NaCl, and 0.1% (w/v) BSA Enriched MAbs (IgG) (see Immunoaffinity Chromatography) Protein A-gold (10 nm; Biocell) diluted 1:10 in Buffer A Flat-topped shaker Reynold's lead citrate 2% (w/v) aqueous uranyl acetate
Procedures FOR CYTOSOLIC ANTIGENS
An equal volume of 1% (v/v) glutaraldehyde in PBS is added to a 500-ml mycoplasma culture and the mixture is incubated (4°C, 30 minutes). After centrifugation (25,000 g, 4°C, 20 minutes), the pellet is dehydrated through an ascending ethanol series: 30% (v/v), 50% (v/v), and 70% (v/v); the pellet is kept in each concentration for 15 minutes. In a suitable container, the pellet is suspended in 4 ml L.R. White resin: 70% (v/v) ethanol, 1:3 (v/v), and rotated (20°C, 1 hour). This is followed by rotation of the pellet overnight in 4 ml L.R. White resin alone. After rinsing the pellet (3x) with L.R. White resin, the final pellet is transferred to Beem capsules which are filled with L.R. White resin prior to polymerization at 50°C in the absence of air (achieved by placing an inverted capsule lid on top of the filled capsule in a vacuum oven). Thin sections are cut (700 A) on an ultramicrotome, collected on water, and transferred to copper grids which are then incubated (4°C, 18 hours) on top of droplets of enriched MAb (diluted 1:10 in Buffer A). After thorough washing of the grids with Buffer A, they are incubated (20°C, 1 hour) on 20-|JL1 droplets of diluted protein A-gold. After thorough washing with Buffer A, sections on the grids are stained with Reynold's lead citrate (5 minutes), washed with distilled water (30 seconds), and then counterstained with 2% (w/v) aqueous uranyl acetate (5 minutes). After washing and air drying, the grids are examined on an electron microscope. FOR SURFACE-EXPRESSED ANTIGENS
Since epitopes of membrane antigens are often lost on glutaraldehyde fixation, it is usually necessary to allow interaction among the cells, MAb, and protein A gold prior to fixation. If antigens are expressed mainly or solely on the interior
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David Thirkell and Bernard L. Precious
Fig. 5. Localization of the 16/17-kDa antigens of U. urealyticum. The mycoplasmas were treated with PBS-N to Increase permeability, probed with MAb UU8/39 and protein A-gold, fixed in 2.5% (v/v) glutaraldehyde in PBS, and embedded in L.R. White. Bar: 0.2 jjim (from Myles eta/., 1991).
surface of the membrane, it may also be necessary to increase the permeability of the mycoplasma cells to allow good access for the MAbs. Mycoplasma cells from a 500-ml culture are pelleted (25,000 g, 4°C, 20 minutes). With no treatment to increase permeability, the pellet is suspended in 2 ml PBS containing 0.2 ml enriched MAb (IgG). To increase permeability, the pellet is suspended in 2 ml PBS containing 0.5% (v/v) Nonidet P-40 and 0.2 ml of enriched MAb (IgG). The suspensions are incubated on ice (2 hours), the cells are pelleted and washed (3x) with Buffer A, and the final pellet is suspended in 1 ml of a 1:10 dilution of protein A-gold in Buffer A. This suspension is incubated, with shaking (20°C, 1 hour). After centrifugation, the pellet is washed (3x) with PBS and then fixed in 2.5% (v/v) glutaraldehyde in PBS (4°C, 15 minutes). After centrifugation, the pellet is dehydrated through an ethanol series and embedded in L.R. White resin as described earlier. Thin sections are cut, stained with lead citrate, counterstained with uranyl acetate, and examined on an electron microscope as before. A typical result for the 16/17-kDa surfaceexpressed antigens of U. urealyticum using the increased permeability method is shown in Fig. 5. Discussion
The MAb binds to its epitope and this is then complexed with protein A-gold. Being electron opaque, the gold particles are seen on the electron microscope as
B8 Immunoblots and Immunobinding
167
black dots allowing interpretation of the cellular location of the antigen. The antigen-antibody reaction must be stable to all procedures.
References Bordier, C. (1981). Phase separation of integral membrane proteins in Triton X-114 solution. J. Biol. Chem. 256, 1604-1607. Coligan, J. E., Kruisbeek, A. M., Margulies, D. H., Shevach, E. M., and Strober, W., eds. (1994). "Current Protocols in Immunology." Wiley, New York. Kessler, S. W. (1975). Rapid isolation of antigens from cells with a staphylococcal protein-A antibody absorbent: Parameters of the interaction of antibody-antigen complexes with proteinA. J. Immunol 115, 1617-1627. Laskey, R. A., and Mills, A. D. (1975). Quantitative film detection of ^H and '"^C in polyacrylamide gels by fluorography. Eur. J. Biiochem. 56, 335-341. Mouches, C , and Bove, J. M. (1983). Electrophoretic characterization of mycoplasma proteins. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 241-255. Academic Press, New York. Myles, A. D., Russell, W. C , Davidson, I., and Thirkell, D. (1991). Ultrastructure of Ureaplasma urealyticum, serotype 8 and the use of immunogold to confirm the localisation of urease and other antigens. FEMS Microbiol Lett. 80, 19-22. Precious, B. L., Thirkell, D., and Russell, W. C. (1987). Preliminary characterization of the urease and a 96 kDa surface-expressed polypeptide of Ureaplasma urealyticum. J. Gen. Microbiol. 133, 2659-2670. Randall, R. E, Young, D. P., Goswami, K. K. A., and Russell, W. C. (1987). Isolation and characterization of monoclonal antibodies to Simian virus 5 and their use in revealing antigenic differences between human, canine and simian isolates. J. Gen. Virol. 68, 2769-2780. Thirkell, D., Spooner, R. K., Jones, G. E., and Russell, W. C. (1990a). Polypeptide and antigenic variability among strains of Mycoplasma ovipneumoniae demonstrated by SDS-PAGE and immunoblotting. Vet. Microbiol 21, 241-254. Thirkell, D., Myles, A. D., and Precious, B. L. (1990b). Characterization of urease and surfaceexpressed antigens of Ureaplasma urealyticum. Zentralbl Bakteriol, Suppl 20, 546-554.
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B9 DIFFERENTIATION OF Mycoplasma genitalium FROM Mycoplasma pneumoniae BY IMMUNOFLUORESCENCE Joseph G. Tully
General Introduction Strains of Mycoplasma genitalium were first isolated from the urethra of men with nongonoccocal urethritis in 1980. Although the organism was clearly established in early studies as a distinct species, it was found to share a number of important biologic and serologic properties with strains of M. pneumoniae (Tully et aL, 1981; Taylor-Robinson et al., 1983; Lind et al, 1984). Thus, the two mycoplasmas have similar organized attachment structures (Tully et aL, 1983) and exhibit sequence homology between their adhesin genes (Dallo et aL, 1989), as well as sharing common epitopes among their adhesin and membrane proteins (Morrison-Plummer et aL, 1987), and they contain similar membrane glycolipids (Taylor-Robinson et aL, 1983; Lind et aL, 1984). Since the respiratory tract was obviously established as the primary site of M. pneumoniae colonization, the interactions with M. genitalium were not thought at the time to complicate the delineation of the role of M. genitalium in human genital tract infections. However, the subsequent discovery of M. genitaliumlM. pneumoniae mixtures in nasopharyngeal throat specimens of patients with acute respiratory disease (Baseman et aL, 1988) not only contributed new concepts about the host distribution of M. genitalium, but prompted important questions about the poten169 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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Joseph G. Tully
tial pathogenicity of the organism and its interrelationship with M. pneumoniae (Tully and Baseman, 1991). A synovial fluid isolate from a patient with polyarthritis following an acute respiratory disease was found to contain both M. genitalium and M. pneumoniae (Tully et al, 1995). With the use of conventional laboratory serologic techniques for mollicutes (growth inhibition and agar plate immunofluorescence tests), each Mycoplasma species was identified within the mixed culture, and individual strains were recovered and their identity was reconfirmed. The applications of these diagnostic techniques are described in detail here. Rapid developments have occurred in the application of DNA amplification procedures (polymerase chain reaction; PCR) to the detection of M. genitalium in clinical specimens, as well as the ability of selected primers to differentiate this organism from M. pneumoniae (de Barbeyrac et al, 1993; Homer et al., 1993; Jensen et al., 1991; Palmer et al, 1991a,b) (see also Section A, this volume). These procedures will obviously become increasingly important in the identification of such fastidious and difficult to cultivate mollicutes as M. genitalium. Likewise, immunoblotting, with mycoplasma cell suspensions and monoclonal antibodies specific either to the adhesin protein of M. pneumoniae (the 168-kDa PI) or M. genitalium (the 140-kDa MgPa), has also successfully been applied to the identification of mixed populations of the two organisms (Baseman ^f«/., 1988).
Materials M. genitalium and M. pneumoniae type or representative strains, grown in SP-4 broth medium (see Chapter A2 in Vol. I.) Agar plates, SP-4 formulation Polyclonal antiserum to type strains of M. genitalium and M. pneumoniae, preferably prepared in rabbits Fluorescein-conjugated antiserum to type strains of M. genitalium andM. pneumoniae. Freeze at -20°C. [If conjugated antiserum is not available, polyclonal rabbit antiserum can be used in an indirect immunofluorescence test, employing fluorescein-conjugated, anti-rabbit (IgG, IgA, IgM) goat serum (Cappel No. 55652, Organon Teknika Corp., Durham, N C ] Phosphate-buffered saHne (PBS), pH 7.8 A fluorescence microscope system, such as the Zeiss standard fluorescence model (with quartz halogen lamp for both incident and transmitted illumination and Zeiss 48-77-05 filter system) (Gardella et al, 1983) Screw-cap glass vials (1-dram or about 4-ml size), sterile Glass or plastic petri dishes (90-mm diameter)
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GasPak incubation system (BBL Microbiology Systems, Cockeysville, MD), with carbon dioxide generation envelopes
Procedure CONTROL DIRECT IMMUNOFLUORESCENCE TESTS WITH ESTABLISHED MYCOPLASMA STRAINS
1. Prior to agar plate immunofluorescence tests on the candidate culture, it is necessary to establish a number of critical parameters related to the responses of established strains of the two mycoplasmas, to variations in the potency of the individual conjugates, to the selection of appropriate dilutions of these conjugates, and to possible variations in the results obtained with different fluorescence microscope systems. 2. Appropriate dilutions of SP-4 broth cultures of the type or representative strains of M. genitalium and M. pneumoniae are plated to individual SP-4 agar and agar plates incubated in the GasPak system in a carbon dioxide atmosphere. Plates selected for testing should contain about 100-200 individual and wellspaced colonies. 3. Thaw frozen fluorescein-conjugated antiserum to each mycoplasma and prepare a series of eight two-fold dilutions (starting a 1:8 and ending at 1:1024) in a final volume of about 2 ml PBS. 4. With a flamed spatula or wire loop, cut eight individual 1-cm-square agar pieces from the plate of M. genitalium colonies and place each piece on an individual glass microscope slide. Label each slide with the appropriate conjugate dilution to be tested. To prevent drying, place slides in a covered petri dish containing two glass rods. Place a filter paper circle in the bottom of the dish and wet with water. 5. With individual capillary pipettes, add 1-2 drops of the appropriate conjugate dilution to the surface of each agar piece, cover, and incubate for 15 minutes at room temperature. Examine each slide and add further conjugate to each agar piece, if necessary, and continue incubation for an additional 15 minutes. 6. At the end of the incubation, gently wash the conjugate from the agar piece with PBS and allow agar to dry for 5-10 minutes. The colonies are embedded in the agar and should not, under most circumstances, become removed by the washing procedure. 7. Examine each agar piece in the fluorescence microscope, using a magnification of about 160x (16x objective and 10x eyepiece). Colonies should first be located on the agar with transmitted white light and then with incident ultraviolet light from the quartz halogen lamp. 8. M. genitalium colonies treated with low dilutions (1:8) of homologous
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Joseph G. Tully TABLE I EPI-IMMUNOFLUORESCENCE TESTS FOR DIFFERENTIATION OF AGAR COLONIES
OF Mycoplasma genitalium FROM Mycoplasma pneumoniae"^ Fluorescence of agar colonies when treated with indicated dilutions of each specific conjugate M. genitalium[
M. pneumoniae 1:128
1:32
1:256
1:1024
3+ 4+ 4+
Neg 2+
Neg
Neg Neg
1+
+
3+
Neg
Neg
Neg
Neg
4+
2+ to 3+
1+
Antigens (agar colonies)
1:16
1:64
M. genitalium M. pneumoniae Strain UTMB-10 (primary isolate) Strain UTMB-lOG (cloned M. genitalium) Strain UTMB-lOP (cloned M. pneumoniae)
4+ 3+ 3+
1+ Neg
4+ 4+
+
2+
"Reproduced from Tully et al., 1995, with permission of the American Society of Microbiology.
conjugate should exhibit strong fluorescence (4-I-), and fluorescence should fade as agar colonies are exposed to more dilute conjugates. The end point dilution of each conjugate is that dilution which still produces a 1+ fluoresence, whereas the working dilution is that which yields a 3-4+ fluorescence. Record the end point and working dilution of the M. genitalium conjugate tested. Repeat this procedure with M. genitalium colonies treated with the M. pneumoniae conjugate. 9. In a similar manner, test M. pneumoniae colonies with a series of M. pneumoniae conjugate dilutions, recording the end point and working dilutions of the specific conjugate. As described earlier, the M. pneumoniae colonies are then tested with the M. genitalium conjugate, and similar end point and working dilutions are recorded. 10. These control tests should establish the dilution of each conjugate which will clearly induce fluorescence in colonies of the homologous organism but will not yield fluorescence in colonies of the other mycoplasma. Table I shows the results with established strains of M. genitalium andM. pneumoniae and several selected dilutions of two conjugated antisera.
Cultivation of Test Isolate in Broth 1. The isolate to be tested for mixed infection should be examined immediately after primary isolation from the host and, if possible, with no more than two or three passages on artificial media. The isolate is then grown on SP-4
B9 Differentiation of M. genitalium
1 73
broth, preferably in a plastic T-25 tissue culture flask containing 10 ml of broth. The culture flask is incubated horizontally at 37°C until there is evidence of attached growth to the plastic surface under the broth layer, as well as some slight color change (toward yellow) in the phenol red indicator in the medium. 2. Pour the supernatant fluid into a discard container for decontamination and add 3-4 ml of fresh SP-4 broth to the flask. Using a sterile tissue cell scraper or rubber policeman, rub the plastic surface to remove the adherent mycoplasmas. Place the flask in an upright position for a few minutes to drain the fluid and then remove the cell suspension with a sterile pipette. Divide the suspension into about two equal volumes in small screw-cap glass vials (4-ml size) and freeze at -70°C.
Preparation and Inoculation of Agar Plates of Test Isolate
1. Prepare a series of five glass vials containing 1:8 ml of SP-4 broth. Thaw a vial of the frozen mycoplasma suspension to be tested and add 0.2 ml to the first vial in the series (final 1:10 dilution). Continue serial 10-fold dilutions of the inoculum by passage of 0.2-ml volumes until the culture is diluted through the last vial (1:10-^). Refreeze the remaining cell suspension material at -70°C. 2. Plate 0.2-ml volume from each individual dilution vial onto two fresh SP-4 agar plates. Mark on the plates the respective dilution factor of the mycoplasma cell suspension. After the inoculum on each agar plate has dried, place the plates in the GasPak jar and incubate in a carbon dioxide atmosphere at 37°C. 3. At 4- to 5-day intervals, remove plates from the incubator and examine microscopically for the appearance and number of mycoplasma colonies on the agar surface. Plates selected for immunofluorescence testing should have numerous but well-spaced individual colonies. Plates containing confluent growth or those with only 10-20 colonies should not be used for subsequent tests. 4. If the initial dilution series yields an unsatisfactory agar colony population for testing, the number of colonies on the plates should indicate whether the culture dilution should be increased or decreased. The frozen mycoplasma suspension can then be thawed, diluted, and again used to prepare a second series of plates, with two agar plates for each dilution.
Immunofluorescence Tests on Agar Colonies of Test Isolate
1. Wet the agar colonies by adding about 5 ml of PBS to each of the two agar plates containing appropriate numbers of colonies of the test organism. Incubate for 5-10 minutes, and then discard the fluids in a container of disinfectant. Tip the dish slightly to drain off excess fluid and remove any remaining fluid with a capillary pipette.
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2. Add about 1-2 ml of a freshly prepared dilution of the selected conjugate of M. genitalium to one plate and an appropriate dilution of the M. pneumoniae conjugate to the second plate. Mark plates with respective conjugate and dilution. Incubate plates for 30 minutes at room temperature. Pour off conjugate into disinfectant solution, wash plates several times with about 5 ml of PBS, and discard wash fluid in disinfectant. Tip plates slightly to drain excess fluid and remove with a capillary pipette. 3. Examine each agar plate in the fluorescence microscope and observe the number and size of colonies showing fluorescence with either conjugate. M. genitalium colonies in a mixture with M. pneumoniae will usually show a smaller size colony, and the numbers may be only 1-5% of the total number of M. pneumoniae colonies.
Separation/Cloning of Isolates from M, genitalium/ M. pneumoniae Mixtures 1. Primary clinical isolates exhibiting mixed populations of M. genitalium and M. pneumoniae frequently have low numbers of M. genitalium. Since it is usually very difficult to select and separate the few individual M. genitalium colonies in this type of mixed population, some effort is necessary during early passages of the isolate to shift the population of M. genitalium colonies toward a 50-50 ratio. 2. The earliest available passage of the mixed culture is grown to the logarithmic phase of growth in SP-4 broth. Four dilutions (1:50, 1:100, 1:150, and 1:200) of the broth culture are plated onto SP-4 agar, using about 0.2 ml per plate. The remaining broth culture is frozen at -70°C. After the inoculum has dried, four 6-mm filter paper disks, saturated with polyclonal antiserum to M. pneumoniae, are placed on the surface of each 60-mm SP-4 agar plate. The disks should be located about halfway from the center to the periphery of the plate surface and about an equal distance from each other. The position of the disks should leave a clear zone of about 2.0 to 2.5 cm^ in the center of the plate. The agar plates are then incubated in the GasPak jar as outlined earlier. 3. At 4- to 5-day intervals, the plates are removed from the incubator and examined under low power magnification (60x). Colonies appearing in the center of the agar surface, in the zone where the M. pneumoniae antiserum has diffused, are selected for subculture to broth. At least one of the plates receiving the diluted test organism should show a moderate number of individual organisms growing within the central area. 4. Aseptically remove agar pieces from the central area of as many of the plates as possible, transferring each to individual screw-cap vials containing about 2.5 ml each of SP-4 broth. Incubate the vials at 37°C and examine in 5-10 days for turbidity or pH changes. When such changes occur, plate the broth
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cultures, with appropriate dilutions, to SP-4 agar and incubate plates anaerobically. 5. Agar plates containing colonies selected from the central zone containing M. pneumoniae antiserum should then be tested in the immunofluorescence test. Individual agar pieces are removed to slides and are stained with conjugates to both M. pneumoniae and M. genitalium. These tests should indicate whether significant shifts in the population of M. genitalium have occured and should show the differences in the colony morphology of the two Mycoplasma species. M. genitalium colonies are usually smaller in size than M. pneumoniae colonies on SP-4 agar. 6. The continued selection of agar colony types and immunofluorescence testing with appropriate dilutions of the two conjugates should provide for the eventual isolation of each individual Mycoplasma species in the mixture. Such isolates should then be subjected to conventional filtration cloning techniques (Tully, 1983) and to reconfirmation by immunofluorescence tests with conjugates employed earlier (Table I).
Discussion Although only few isolates of M. genitalium have been made from humans, experience indicates that the organism is extremely fastidious and has an exceedingly slow growth rate. In those instances where the organism has occurred along with M. pneumoniae, it has been obvious that continued passage of the mixture will not favor the survival of M. genitalium. Where the two organisms were found in a synovial fluid isolate, the culture had been passaged about five times. When examined by the differential immunofluorescence test outlined in this chapter, the number of M. genitalium colonies on the initial plating of the synovial isolate was less than 1% of the total number of agar colonies obtained. Detection procedures for M. genitalium, regardless of whether the immunofluorescence or immunoblotting techniques are utilized, should be performed only on early passages of clinical specimens. It should be emphasized that each conjugated antiserum employed in the test must be evaluated in standard immunofluorescence tests against the homologous mycoplasma strain before such conjugates can be applied to potentially mixed cultures. Likewise, if the indirect immunofluorescence method is employed, a checkerboard program must be performed with agar colonies of the representative mycoplasma. In this program, various dilutions of the unlabeled, polyclonal antiserum are compared to various dilutions of the labeled, anti-rabbit conjugate, with the view of selecting the optimum level of conjugate necessary for the mycoplasma antiserum. The eventual delineation of the role of M. genitalium in human disease,
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involving either the urogenital or respiratory tracts, or other tissue sites, will depend on the further application of rapid diagnostic techniques, laboratory isolation, and identification by conventional methodology outlined here, and a serologic analysis of host immune responses by specific and differential techniques.
References Baseman, J. B., Dallo, S. F., Tully, J. G., and Rose, D. L. (1988). Isolation and characterization of Mycoplasma genitalium strains from the human respiratory tract. J. Clin. Microbiol. 26, 22662269. Dallo, S. F., Chavoya, A., Su, C.-J., and Baseman, J. B. (1989). DNA and protein sequence homologies between the adhesins of Mycoplasma genitalium and Mycoplasma pneumoniae. Infect. Immun. 57, 1059-1065. de Barbeyrac, B., Bemet-Poggi, C , Febrer, F., Renaudin, H., Dupon, M., and Bebear, C. (1993). Detection of Mycoplasma pneumoniae and Mycoplasma genitalium in clinical samples by polymerase chain reaction. Clin. Infect. Dis. 17(Suppl. 1), S83-S89. Gardella, R. S., Del Giudice, R. A., and Tully, J. G. (1983). Immunofluorescence. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, pp. 431-439. Academic Press, New York. Homer, P. J., Gilroy, C. B., Thomas, B. J., Naidoo, R. O. M., and Taylor-Robinson, D. (1993). Association of Mycoplasma genitalium with acute non-gonococcal urethritis. Lancet Ml, 582585. Jensen, J. S., Uldum, S. A., S0ndergard-Anderson, J., Vuust, J., and Lind, K. (1991). Polymerase chain reaction for detection of Mycoplasma genitalium. J. Clin. Microbiol. 29, 46-50. Lind, K., Lindhardt, B. O., Schutten, H. J., Blom, J., and Christiansen, C. (1984). Serological cross-reactions bQiwccn Mycoplasma genitalium and Mycoplasma pneumoniae. J. Clin. Microbiol. 20, 1036-1043. Morrison-Plummer, J., Lazzell, A., and Baseman, J. B. (1987). Shared epitopes between Mycoplasma pneumoniae major adhesin protein PI and a 140-kilodalton protein of Mycoplasma genitalium. Infect. Immun. 55, 49-56. Palmer, H. M., Gilroy, C. B., Claydon, E. J., and Taylor-Robinson, D. (1991a). Detection of Mycoplasma genitalium in the genitourinary tract of women by the polymerase chain reaction. Int. J. STD AIDS 2, 261-263. Palmer, H. M., Gilroy, C. B., Furr, P. M., and Taylor-Robinson, D. (1991b). Development and evaluation of the polymerase chain reaction to detect Mycoplasma genitalium. FEMS Microbiol. Lett. 77, 199-204. Taylor-Robinson, D., Furr, P. M., and Tully, J. G. (1983). Serological cross-reactions between Mycoplasma genitalium and M. pneumoniae. Lancet 1, 527. Tully, J. G. (1983). Cloning and filtration techniques for mycoplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, pp. 173-177. Academic Press, New York. Tully, J. G., and Baseman, J. B. (1991). Mycoplasma. Lancet 337, 1296. Tully, J. G., Taylor-Robinson, D., Cole, R. M., and Rose, D. L. (1981). A newly discovered mycoplasma in the human urogenital tract. Lancet 1, 1288-1291. Tully, J. G., Taylor-Robinson, D., Rose, D. L., Cole, R. M., and Bove, J. M. (1983). Mycoplasma
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genitalium, a new species from the human urogenital tract. Int. J. Syst. Bacteriol. 33, 387396. Tully, J. G., Rose, D. L., Baseman, J. B., Dallo, S. F., Lazzell, A. L., and Davis, C. P. (1995). Mycoplasma pneumoniae and Mycoplasma genitalium mixture in synovial fluid isolate. J. Clin. Microbiol. 33, 1851-1855.
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SECTION
C
Antibiotic Sensitivity Testing
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CI INTRODUCTORY REMARKS Christiane Bebear
The absence of a cell wall as a major structural component of moUicutes has significantly restricted the selection of antibiotic drugs that might be utilized against these organisms. In general, all mollicutes are completely insensitive to antibiotic compounds that are directed to inhibition of cell wall synthesis, such as p-lactam antibiotics. In addition, mollicutes are generally resistant to polymyxins, rifampin, and the sulfonamides. Antibiotics exhibiting the most inhibitory activity toward mollicutes, and the most useful in treatment of infections with these organisms, are the tetracyclines, the macrolides and related antibiotics, and some fluoroquinolones. Other antibiotics, such as the aminosides and chloramphenicol, possess less inhibitory activity toward mollicutes and are therefore not very useful in chemotherapy of infections by these organisms. However, some variation in inhibitory response can be observed within specific mollicutes and with certain antibiotics. Innate resistance to erythromycin, but not to all other macrolides, is usually observed with strains of Mycoplasma hominis and M. fermentans, whereas Ureaplasma urealyticum strains are generally resistant to lincomycin. Also, it is necessary to emphasize that the most active antibiotics that are successful in chemotherapeutic treatment of mollicute infections induce a bacteriostatic effect rather than a lethal response on the organism. Antibiotic sensitivity tests on mollicutes have shown a few consistent and well-documented instances where a specific antibiotic has produced a permanent lethal effect against a variety of strains of the organism, especially within drug levels employed in conventional clinical situations. Additional support for these impressions can be derived from difficulties in the consistent eradication of mollicutes from contaminated cell culture infections with various antibiotics, and the inabil181 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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ity of antibiotics to eradicate mollicute infections, either in patients with immune deficiencies or in hosts (such as plants) with no functioning immune system. An in vitro antibiotic sensitivity assay is obviously an important and necessary procedure whenever a mollicute with an established disease relationship to the host is isolated and identified, and when a course of chemotherapy is being considered. Also, one must always consider the possibility that the particular mollicute in question has developed an acquired resistance to the drug of choice, and sensitivity testing can provide information for selection of the most active antibiotic. Acquired resistance to various antibiotics has now been well documented within various moUicutes. However, the occurrence of antibiotic resistance depends a great deal on the particular species of mollicute, the colonization site in the host, and the particular antibiotic. Acquired resistance to macrolides has been reported only in a few isolates of M. pneumoniae, especially those organisms purposely cultivated in the presence of erythromycin or in strains isolated from patients receiving extensive chemotherapy with the drug. Such strains were resistant to several macrolides, including lincomycin and streptogramin B. At this time, the occurrence of M. pneumoniae strains originating from the human respiratory tract that are resistant to tetracyclines or quinolones has not been observed or reported. Currently, antibiotic resistance in mollicutes isolated from the human urogenital tract presents a very different picture. Acquired resistance to tetracycline has been recorded in about 5% of U. urealyticum and M. hominis strains isolated in France. This resistance is apparently mediated through genetic transfer among other urogenital bacteria carrying the tetM resistance determinant and genital tract mollicutes. The occurrence of such resistance is now thought to relate to therapeutic treatment failures, particularly in clinical settings (i.e., nongonococcal urethritis due to U. urealyticum) where tetracycline has been the drug of choice. Acquired resistance to erythromycin has been reported in strains of U. urealyticum, but tests for in vitro susceptibility with this antibiotic are difficult because of the low pH required for growth of the organism and the deleterious effect of acidic environments on the drug. Unlike the standardized protocols for antibiotic sensitivity testing of bacteria, no formalized technique has been developed for mollicutes. A working team of the International Programme on Comparative Mycoplasmology, a component of the International Organization for Mycoplasmology, has been involved in developing some guidelines for such standardization. Some of the requirements for standardization and various techniques that have been employed in antibiotic sensitivity tests with mollicutes are reviewed in C2 of this section. Chapter C3 describes in some detail currently recognized procedures for determination of minimal inhibitory concentration of antimicrobial agents acting on mollicutes. As mentioned earlier, the inability of most antibiotic compounds to provide a
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lethal effect on mollicutes has prompted the development of laboratory procedures that more adequately define the amount of the drug that provides a bacteriostatic effect versus a bactericidal response. Chapter C4 outlines procedures that have been successfully employed in measuring bactericidal responses in mollicutes.
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C2 PROBLEMS AND OPPORTUNITIES IN SUSCEPTIBILITY TESTING OF MOLLICUTES George E. Kenny
Introduction The major purpose in susceptibility testing of mycoplasmas is to determine the in vitro susceptibility of a given moUicute species to new and older antimicrobial agents in an effort to identify agents which might have clinical utility in the treatment of mycoplasmal infections. Overall, the methodology thus far developed has not achieved consensus because the susceptibilities reported have varied widely in the literature and the range of susceptibilities for a given species has been extremely wide (Roberts, 1992), suggesting a need for the development of methodology. This chapter presents some concepts which may provide a basis for perfecting methods for mycoplasmal susceptibility testing, an area which should provide excellent opportunities for research. Methods for susceptibility testing of bacteria are well established for conventional pathogens, such as Staphylococcus aureus, which grow rapidly and form colonies within 24 hours. The medium to be used, inoculum size, and incubation period have been defined for two methods: broth dilution where the end point is turbidometric (Amsterdam, 1991) and agar dilution where the end point is colony formation. The third method, disk diffusion, is not applicable to slow-growing organisms because the agent will equilibrate across the plate before growth. Clinical laboratories follow exact protocols with specific control organisms. In contrast, susceptibility testing methods for organisms which grow more slowly, 185 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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such as mycobacteria, are less well established, are still in the developmental phase, and are not so widely used.
Problems in Susceptibility Testing The specific problems include: (1) inoculum size, (2) incubation period, (3) medium and its pH, and (4) phase of growth. A major challenge in interpretation of results is posed by (5) mycoplasmacidal testing, but this challenge provides a fertile field for future research (see Chapter C4, this volume). Inoculum Size The failure of mycoplasmas to produce measurable turbidity is a major problem for determining inoculum size in contrast to conventional bacteria where the inoculum can be determined turbidometrically with very small error. As a consequence, several dilutions of culture must be tested in order to obtain a defined inoculum. Quantitation of the inoculum has frequently been carried out by colorimetric assays where only a single well or tube is used per 10-fold dilution. Such methods can produce 10-fold differences in inoculum size (Koch, 1981), and this variation contributes to the large ranges reported for minimum inhibitory concentration (MIC) values reported for isolates of a single species. For most agents, susceptible strains show quite narrow ranges of susceptibility to quinolones and tetracyclines (Kenny and Cartwright, 1993a, 1994). The inoculum size needs to be reproduced within 2-fold in order to provide for reproducible results in the broth dilution method (Kenny and Cartwright, 1993b). Quantitative methods such as plate count, determination of 50% end points, or most probable numbers using multiple replicates, need to be used to quantitate inocula (Koch, 1981). Luminometry (see Chapter C3 in this volume) may provide a means of standardization of inocula. Incubation Period The reporting of an initial and a final MIC does not directly answer the question of whether an organism is susceptible or not. The incubation period for the assay could probably be set at twice the length of time (or some other multiple) it takes a culture of the particular species to form colonies or produce a measurable end product. This will be longer for organisms which grow slowly, such as Mycoplasma pneumoniae (6-hour generation time), compared with organisms which grow rapidly, such as M. hominis (1.5-hr generation time).
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Medium It is not possible to standardize on a single medium since growth requirements of mycoplasmas differ. The addition of large concentrations of cations to medium for ureaplasmas could cause serious problems for certain antimicrobials. Such cations are just as effective for visualizing colonies when added at the end of the incubation period. The pH value of the medium causes a significant difference in testing macrolides (Kenny and Cartwright, 1993b) and a lesser, though significant, difference in testing tetracyclines (Kenny and Cartwright, 1994). For mycoplasmas, the pH should be standardized to a physiological value of pH 7.3 rather than the usually used pH 7.6-7.8 value. The pH should be measured with a surface electrode during incubation. Ureaplasmas cause problems because they grow so poorly at neutral pH that testing is impossible. However, testing at or about pH 6.3 will be relevant to urinary tract infections and possibly to vaginal colonization. Phase of Growth For initial surveys with new antibiotics, it is important that mycoplasmas be in the logarithmic phase of growth since these conditions are commonly used for conventional bacteria. Thus, frozen and stored cultures should not be used in such evaluations. However, the use of nongrowing cultures as inocula should be studied because it has been shown that bacteria which are growing slowly because of deficient media show static effects with known bactericidal agents (Eng etaL, 1991). Bactericidal Testing The conventional method for determining bactericidal effects with bacteria is the time-kill method where a mixture of bacteria and antibiotic is sampled periodically and plate counted to determine the survival of the bacteria. A reduction in the plate count of 99.9% or greater in 24 hours or less is taken as evidence of bactericidal effects for fast-growing bacteria. In mycoplasmas, incubation periods have varied from several to many days. As a consequence, wellestablished bacteriostatic agents such as tetracyclines and macrolides have been reported to be mycoplasmacidal in some but not all reports. The time-kill method needs to be studied in more detail to determine the kinetics of killing mycoplasmas by antibiotics. Mycoplasmas pose unique problems since many species show a steep death phase immediately after peak growth, which may confuse killing by the agent with death from "old age" (Taylor-Robinson and Furr, 1982; see also Chapter C4, this volume).
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References Amsterdam, D. (1991). Susceptibility testing of antimicrobials in liquid media. In "Antibiotics in Laboratory Medicine" (V. Lorian, ed), pp. 53-119. Williams & Wilkins, Baltimore. Eng, R. H. K., Padberg, F. T., Smith, S. M., Tan, E. N., and Cherubin, C. E. (1991). Bactericidal effects of antibiotics on slowly growing and nongrowing bacteria. Antimicrob. Agents Chemother. 35, 1824-1828. Kenny, G. E., and Cartwright, F. D. (1993a). Susceptibilities of Mycoplasma hominis. Mycoplasma pneumoniae, and Ureaplasma urealyticum to a new quinolone, OPC-17116. Antimicrob. Agents Chemother. 37, 1726-1727. Kenny, G. E., and Cartwright, F. D. (1993b). The effect of pH, inoculum size, and incubation time on the susceptibility of Ureaplasma urealyticum to erythromycin in vitro. Clin. Infect. Dis. 17(Suppl. 1), S215-S218. Kenny, G. E., and Cartwright, F. D. (1994). Susceptibilities of Mycoplasma hominis. Mycoplasma pneumoniae, and Ureaplasma urealyticum to new glycylcyclines in comparison with those to older tetracyclines. Antimicrob. Agents Chemother. 38, 2628-2632. Koch, A. L. (1981). Growth measurement. In "Manual of Methods for General Bacteriology" (P. Gerhardt, R. G. E. Murray, R. N. Costilow, E. W. Nester, W. A. Wood, N. R. Krieg, and G. B. Phillips, eds.), pp. 179-207. Am. Soc. Microbiol., Washington, DC. Roberts, M. C. (1992). Antibiotic resistance. In "Mycoplasmas: Molecular Biology and Pathogenesis" (J. Maniloff, R. N. McElhaney, L. R. Finch, and J. B. Baseman, eds.), pp. 513-523. Am. Soc. Microbiol., Washington, DC. Taylor-Robinson, D., and Furr, P. M. (1982). The static effect of rosaramicin on Ureaplasma urealyticum and the development of antibiotic resistance. J. Antimicrob. Chemother. 10, 185191.
C3 DETERMINATION OF MINIMAL INHIBITORY CONCENTRATION Christiane Bebear and Janet A. Robertson
Introduction The minimal inhibitory concentration (MIC) is the highest dilution of an antimicrobial agent that inhibits the growth of a particular microorganism in the test period. MIC tests are well established in bacteriology and constitute the most common means of determining mycoplasmal susceptibility to antimicrobial agents. Because a well-designed and well-executed MIC test should yield reproducible data for a given strain wherever it is performed, rigorous attention to standardization is required. The basic principles of antimicrobial susceptibility testing of bacteria may be found in a comprehensive clinical laboratory manual (Balows et al, 1991). Difficulties associated with testing mycoplasmas have been presented (see Chapter C2 in this volume). Erroneously high MICs have been associated specifically with the relatively long generation times of mycoplasmas, with antibiotic binding by the serum required for growth media, and with the deleterious effects of pH and cation concentrations on the activity of certain agents. MIC tests can be used to determine mycoplasmal responses to potentially inhibitory agents other than antibiotics (Robertson et al., 1988; Ostashewski et al., 1993).
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Materials Media Modified Edward broth for Mycoplasma contains heart infusion broth (1.7%, w/v), fresh yeast extract (2.5%, v/v), horse serum (20%, v/v), and phenol red indicator (0.001%, w/v). Broth for glycolytic species contains added glucose (~0.1-0.5%, w/v; final broth pH ~7.4-7.6), whereas that for argininolytic species contains added arginine (<0.5%, w/v; final broth pH ~7.2-7.4). Optimally, media for the ureolytic Ureaplasma species contain about 0.025-0.1% added urea and are adjusted to a low pH (6.0-6.5). SP4 medium may be used for fastidious mollicutes including Spiroplasma species (see Chapter A2 in Vol. I). Medium is usually solidified by ~ 1% purified agar. Solid media for ureaplasmas usually contain a buffer and may include a CaCl2 indicator system to accentuate and identify, respectively, the tiny colonies of this genus. Media formulations for specific mollicutes species are available elsehwere (Razin and Tully, 1983). Growth medium, appropriate for the test species, should be brought to room temperature for inoculation so that, on incubation, the lag phase is not prolonged. Antibiotic or Chemical Agents Agents should be of known chemical purity; antibiotic-impregnated disks are not a suitable source of agent. Storage requirements, solubility characteristics, pH for optimal activity, and half-lives under test conditions can be obtained from a comprehensive bacteriology laboratory manual (Balows etal., 1991), a recent chemical index, or the manufacturer. If agent stability is unknown, use freshly made solutions. If aminoglycosides are to be tested, medium cation concentrations should be appropriate.The final concentrations of antibiotic should encompass the achievable therapeutic levels of the agent. For antibiotics, the convention is to express MIC values as doublings or halvings of 1 |xg/ml. However, concentrations of other potentially inhibitory agents are more usefully expressed on a molar basis. Organisms An appropriate inoculum should provide the MIC in the minimum time without metabolites compromising accuracy. Inocula should be prepared from the highly responsive cells of cultures in exponential growth. At near maximum growth the number of viable cells per ml may be estimated at —10^ for ureaplasmas or ~10^-109 for most large-colony mycoplasmas. Cell numbers are usually expressed as color-changing units (CCU) or colony-forming units (CFU) for broth or agar medium, respectively. If the culture titer is not known, a CCU
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or CFU count is made of the test culture, MICs are performed on several dilutions of it, and the proper MIC is selected when the titer has been established. Equipment
Desiccators at appropriate temperatures for antibiotic storage Balance for weighing antibiotics or other agents BROTH-BASED TESTS
Microtiter methods require recently calibrated multichannel micropipettors or microdilutors; sterile, disposable tips; microtiter plates; sealing tape; needle or venting apparatus; and sheets for recording results. The tube method requires no specialized equipment, only sterile serological and Pasteur pipettes, rubber teat, and test tubes. AGAR-BASED TESTS
Plates with agar, multipoint replicator, micropipettor, or a calibrated loop. A template for locating inocula and a sheet for recording results are helpful. Incubation of all but obligately aerobic species requires "anaerobic" jars and either analyzed gas or gas generators. A microscope, preferable inverted, is required for examination of plates. ATP-DEPENDENT LUMINOMETRY See Chapter A5 in Vol. I. COMMERCIAL KITS Follow manufacturer's instructions.
Procedures Choice of Test Type
The choice of test procedure is based on such factors as the number of strains to be tested, their growth response in liquid and solid media, generation time of the species, and the half-life of the agent under examination. When small numbers of strains are to be tested or when the minimal bactericidal concentration (see Chapter C4, this volume) is desired, broth methods are more convenient and more economical. General times of moUicutes range from 60 to 90 minutes for ureaplasmas from humans and to 8 hours for M. pneumoniae and 10 hours for the
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porcine species M. hyopneumoniae. When generation times are exceptionally long, evaporation is a problem and necessitates using larger volumes of broth and tube tests. The half-life of antibiotics varies greatly. ATP luminometry (see later) should be considered when a long generation time or a very short antibiotic halflife is a factor. MIC tests and controls are performed in duplicate and incubated under the conditions usually required for the growth of the test species. Controls
Mandatory controls are required for medium, medium containing the highest concentration of agent used in the test, and the growth of each test and control strain in medium alone. Normally, only the last of those controls should change color. When controls misperform, tests are repeated. The effect of the medium on the activity of a new agent is determined by comparing parallel MICs of a reference bacterium of known, low MIC that have been conducted both in standard medium (e.g., Mueller-Hinton) and in mycoplasma medium. When available, strains of the species under examination, but of known susceptibility/ resistance, should be included as controls. These strains can be maintained for years if the broth cultures are stored with —10% sterile glycerol at <-60°C. Reading and Interpretation of Tests
Cultures are incubated until the mycoplasma control broths change color or colonies develop on agar. Presumptive readings (initial MICs) are made then; final MICs are made at a specified time, usually 24 hours later. When the results of duplicate tests differ by one doubling dilution, the more conservative reading is used. When results differ by more than one dilution, the methodology, especially the tools used to prepare the dilutions, should be examined and the test repeated. MICs are interpreted according to concentrations of the test agent attainable in blood, urine, or other body fluid. Broth-Based Tests
Doubling dilutions of potentially inhibitory agents are tested against a standardized inoculum; substrate degradation alters the color of a pH indicator in the medium, signaling growth. A tube method is convenient for testing small numbers of strains or for screening strains for possible resistance. Miniaturized, microtiter versions are the least demanding and most commonly used method of MIC determination. They are efficient for testing large numbers of strains and/or multiple antibiotics. The following protocol uses lOO-jxl capacity, multichannel micropipettors (Bebear et al., 1985); the protocol has been adapted from the 25-|xl microdiluters commonly used in virology.
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MICROTITER METHOD
All tests and controls are set up in duplicate. 1. Add 100 |xl broth to wells 2-12 of each horizontal row of a 96-well microtiter plate. 2. Add 100 |xl of antibiotic-containing broth to wells 1 and 2 of each row. This broth contains twice the highest concentration of antibiotic to be tested. 3. Make doubling dilutions from wells 2 through 12. 4. Add 100 |JL1 of inoculum in fresh broth to each well. Final inoculum concentrations are in the range of 10"^-10^ viable cells per ml. Thus a 10'^ CCU/ml culture of Ureaplasma would be diluted about 1/100 and a lO^-lO^ CCU/ml culture of M. hominis at least 1/1000. 5. Mandatory controls are medium control, 200 jxl broth; antibiotic control, 100 |xl of antibiotic-containing broth and 100 |JL1 broth; organism controls for each test/control strain, 100 \x\ of broth and 100 jxl inoculum. Depending on the solvent required to prepare the stock solution of antibiotic, a solvent control may be included. 6. Seal plates with tape, vent wells, and incubate plate in a humidified cabinet at an appropriate incubation temperature and gaseous environment until the organism controls change color. Record presumptive MICs. Usually, final MICs are taken after 24 hours of further incubation. Presumptive readings, of Ureaplasma and M. hominis are usually available at 16-24 hours and at 36-48 hours, respectively; many other moUicute species will require longer incubation. POTENTIAL PROBLEMS
If plates are examined outside of the incubator and condensation obscures color changes, read wells from the underside of the plate. The horizontal spread of gaseous end products can affect color changes in sterile wells. This can result from use of plastic microtiter covers, moisture interfering with a complete seal, or failing to vent the sealed wells, and is most marked when high substrate concentrations are used. A color change in a broth control indicates contamination. Most mollicutes produce neither turbidity nor a pellet, even by the decline phase; either is indicative of contamination. Discoloration of the antibiotic controls suggests agent deterioration; usually this effect is recognizable as it decreases with antibiotic dilution. If an organism control did not change color within the expected incubation period, retest that strain along with the usual controls. TUBE METHOD
A 10-fold increase in the previously mentioned volumes will convert a microtiter method to a tube method with a final volume of 2 ml. One drop from a
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Pasteur pipette (—25 jxl) of an undiluted or a 1/10 dilution of a culture of ureaplasma or a 1/1000-1/10,000 dilution of a culture of a large colony mycoplasma should provide an appropriate inoculum. Include controls with each test. Agar-Based Tests
For organisms that do not grow well in broth but form colonies on agar (e.g., the avian species, Mycoplasma meleagridis), MICs can be determined on solid medium. Because of the relatively slow growth rate of mollicutes, the antimicrobial agent is incorporated directly into the agar medium rather than waiting for diffusion from paper disks. As one plate is required for each concentration of every antibiotic to be tested, the method is most practical when many strains are to be tested. 1. Agar plates are inoculated with a multipoint replicator, a micropipettor, or a calibrated loop. The goal is to obtain 30-300 colonies per spot of inoculum on the mycoplasma control plate. However, because all organisms in the inoculum may not produce colonies on agar, the equivalent of about 10 |xl of a 1/10 dilution of a M. pneumoniae culture or a 1/100 dilution of a culture of M. hominis may yield the desired result. Because the size of moUicute colonies is inversely related to colony number, heavy inocula can completely mask the presence of growth. Because Ureaplasma colonies are exceptionally small, several dilutions of culture may be used as inocula for multiple tests. The inoculum giving rise to the appropriate number of colonies on the control plate can be selected after incubation. 2. After incubation, inoculum spots on the agar surface are examined using low power microscopy. The MIC is the lowest concentration of agent which completely prevents colony formation when the requisite 30-300 colonies can be detected on the antibiotic-free control plates. This usually occurs at 2 days for M. hominis, at 2-4 days for U. urealyticum, and at 5 or more days for other species of interest in human medicine. POTENTIAL PROBLEMS
These tests depend on the detection of colonies. If an inoculum contains too many cells or if the nutrient or physical conditions of culture are deficient, recognizable colonies may not form. Thus the quality of the test medium is of critical importance. First, the agar or agarose should be from a single lot that has been shown to be noninhibitory to the test species and of a gel strength that allows easily recognizable colonies to develop. Second, before the addition of heat-sensitive medium supplements and antibiotics to the basal medium, the temperature of the molten agar should be ascertained directly rather than deduced from the set temperature of the water bath. Third, as antibiotics deteriorate over
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time, media must be used when fresh. Finally, agar medium is more subject than broth medium to batch-to-batch variation in performance. Nevertheless, agarbased MIC tests give consistent results in the hands of experienced workers (Kenny et aL, 1989) examining species of well-established growth characteristics. Incubation times may be slightly longer and the MICs slightly higher than for tests conducted in broth media (Waites et al., 1991). ATP'Dependent Luminometry (see Chapter A5 in Vol. I) This method is suitable for determining MICs of species that are highly fastidious or have long generation times. It is also useful for testing agents with a short half-life. We suggest adding the agent to early logarithmic phase cultures and comparing subsequent growth with the rate in control cultures. Commercial Kits These kits are available for testing the susceptibility of genital mycoplasmas to antibiotics. Because the number of organisms present in clinical specimens is highly variable, a two-step procedure is required: the first step standardizes the inoculum and the second step tests the response of the organism to the agent. By testing two concentrations of each agent, the mycoplasma can be classified as susceptible, intermediately responsive, or resistant to the antibiotic.
Discussion The responses of some mycoplasmas of humans to selected antibiotic agents are shown in Table I. Variation in strain susceptibility to a particular class of antibiotics is associated primarily with the response of common genital species to tetracyclines (Roberts, 1992). The better known the growth responses of the test species, the easier it is to select the most appropriate method for determining MIC. Then, attention to detail is a requisite for accurate results. The choice of culture medium is extremely important. To promote test accuracy, avoid excessive supplementation with serum or substrate. In regard to MIC testings, serum binding of antibiotics (e.g., tetracyclines) reduces antimicrobial activity, giving erroneously high MICs, whereas the buffering capacity of serum can delay pH indicator changes, giving erroneously low MICs. Because serum has growth-inhibiting effects for certain mollicutes, as well as its more generally recognized growth-promoting effects, increasing the serum supplementation of media will not necessarily provide better growth. Similarly, an ideal substrate concentration would allow
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Christiane Beb^ar and Janet A. Robertson
TABLE I MIC RANGE OF ANTIBIOTICS AGAINST MYCOPLASMAS OF HUMANS''
Mycoplasma pneumoniae
M. genitalium
M. fermentans^
M. hominis
Ureaplasma urealyticum
0.02-0.01 0.02 0.01
<0.01-0.05 <0.01 0.02
0.13-0.26 ND^
0.02-0.05^^ 0.02-1^
0.02-1^ 0.02-K
Erythromycin Roxithromycin Clarithromycin Azithromycin Josamycin
0.03-0.06 <0.01 0.05 <0.01 <0.01-0.02
<0.01 <0.01 <0.01 <0.01 0.02
32-64 ND 16-64 ND 0.2
>128 >16 16-128 4-64 0.05-0.1
0.5-4 0.1-2 0.02-0.2 0.5-4 0.1-1
Lincomycin Clindamycin Pristinamycin
4-8 1-2 0.02-0.05
1-8 0.2-1 <0.01-0.02
0.15-0.2 0.14-0.23 0.02-0.05
0.2-1 0.02-0.05 0.1-0.5
8-256 0.5-16 0.1-1
Pefloxacin Ciprofloxacin Ofloxacin Sparfloxacin
2 1 0.05-1 0.1
ND 2 1-2 0.05-0.1
ND 0.02-0.16 0.1-0.2 <0.01-0.05
0.5-2 0.1-1 0.2-2 <0.01
0.5-8 1-16 0.2-2 0.1-0.5
Antibiotic Doxycycline Minocycline
^Data in micrograms per milliliter. ^From Hayes et al. (1991). ^ Tetracycline-susceptible strains. ^Not determined.
maximum growth and discernible pH changes without effecting the MIC. On the other hand, catabqlism of high concentrations of glucose results in an acid milieu. Acidity markedly reduces macrolide activity and also is believed to limit the growth or viability of certain species (e.g., the avian species, M. synoviae). The effects of a very alkaline milieu on growth or antibiotic activity have not been well defined; this extreme also should be avoided. Inconsistent MICs of clinical isolates could reflect mixed cultures. If each species had distinctive colonial morphology and was present in more or less equal proportions, mixtures might be detected on agar cultures. One can predict, however, that usually one species will predominate and the other will be missed. If mixed cultures are anticipated, selective antibiotic or antiserum activity can be used on primary culture or during inoculum preparation. For instance, the inclusion of lincomycin or clindamycin in medium stops large colony mycoplasmas (e.g., M. hominis) from overgrowing Ureaplasma species. Finally, when determining MICs, convenience should not take precedence over accuracy. If tubed media or microtiter trays are prepared in advance and
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frozen until required, storage conditions are critical and their effects need to be monitored. The practice of using a frozen culture of a previously determined titer as an inoculum also has pitfalls. Viability may be lost during storage or on thawing. Even if this does not occur, the lag phase of thawed cultures is likely to be prolonged over that of a fresh culture and MICs could be falsely elevated. When determining MICs of mollicutes and when methodology is in question, good bacteriologic practice should be the guide.
References Balows, A., Hausler, W. J., Jr., Herrmann, K. L., Isenberg, H., and Shadomy, H. J., eds. (1991). "Manual of Clinical Microbiology," 5th ed. Am. Soc. Microbiol., Washington, DC. Bebear, C , Cantet, P., Renaudin, H., and Quentin, C. (1985). Activite comparee de la minocycline et de la doxycycline sur les mycoplasmas pathogenes pour I'homme. Pathol. Biol. 33, 577580. Hayes, M. M., Wear, D. J., and Lo, S. C. (1991). In vitro antimicrobial susceptibility testing for the newly identified AIDS-associated Mycoplasma. Arch. Pathol. Lab. Med. 115, 464-466. Kenny, G. E., Hooten, T. M., Roberts, M. C , Cartwright, F. D., and Hoyt, J. (1989). Susceptibilities of genital mycoplasmas to the newer quinolones as determined by the agar dilution method. Antimicrob. Agents Chemother. 33, 103-107. Ostashewski, P. M., Houston, S. C , and Robertson, J. A. (1993). In vitro activity of zidovudine, zalcitabine, and didanosine against mycoplasmas. Lancet 342, 1242-1243. Razin, S., and TuUy, J. G., eds. (1983). "Methods in Mycoplasmology," Vol. 1. Academic Press, New York. Roberts, M. C. (1992). Antibiotic resistance in mycoplasmas. In "Mycoplasmas: Molecular Biology and Pathogenesis" (J. Maniloff, R. N. McElhaney, L. R. Finch, and J. B. Baseman, eds.), pp. 512-523. Am. Soc. Microbiol., Washington, DC. Robertson, J. A., Stemke, G. W., MacLellan, S. G., and Taylor, D. E. (1988). Characterization of tetracycline-resistant strains of Ureaplasma urealyticum. J. Antimicrob. Chemother. 21, 319332. Waites, K. B., Figarola, T. A., Schmid, T., Crabb, D. M., Duffy, L. B., and Simecka, J. W. (1991). Comparison of agar versus broth dilution techniques for determining antibiotic susceptibilities of Ureaplasma urealyticum. Diagn. Microbiol. Infect. Dis. 14, 265-271.
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C4 CIDAL ACTIVITY TESTING David Taylor-Robinson
Introduction The antibiotic susceptibilities of moUicutes may be determined in vitro by two methods: the agar dilution method (Roberts, 1992), in which the end point is the prevention of colony development on agar, and a metabolism-inhibition (MI) method (Taylor-Robinson, 1967; Taylor-Robinson and Furr, 1982; Roberts, 1992) using broth medium, in which the antibiotic inhibits growth of the organisms and, hence, their metabolism. The MI method is a simple modification of that used for measuring antibody, the latter being replaced by antibiotic (see Chapter C3, this volume). The MI method is perhaps preferable for most investigators, particularly when testing the susceptibility of ureaplasmas, since color changes in broth medium are easier to demonstrate than colony development. The end point (minimal inhibitory concentration; MIC) in the MI method is the highest dilution of antibiotic at which the color change is inhibited (TaylorRobinson, 1967; Taylor-Robinson and Furr, 1982; Roberts, 1992). While this measurement is satisfactory in most clinical settings, antibiotics active against moUicutes tend not to be cidal, at least in concentrations that can be achieved in vivo, and the MIC value does not reflect the lack of killing activity, nor whether an antibiotic might have such capacity. This chapter outlines the methods used to determine the cidal activity of an antibiotic or the lack of it.
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Materials Media^ Substrates^ and Indicators The basic composition of culture media has been presented elsewhere (Freundt, 1983; Shepard, 1983; Whitcomb, 1983; see also Chapters A2 and A3 in Volume I), as has the nature and choice of substrates to be metabolized and the pH indicators (Taylor-Robinson, 1983).
Mycoplasmas Organisms should be used that multiply easily in the test medium and produce an adequate pH change (or reduction of tetrazolium) in the presence of the specific substrate. A freshly growing broth culture may be used as antigen in the test proper, but the most reproducible results are obtained by dividing a broth culture into aliquots, storing them at -70°C, and using a newly thawed aliquot for each test. One vial should be used for titration of the number of viable organisms and this titration will suffice for all subsequent MI tests for antibiotic sensitivity.
Microtiter Plates^ Micropipettes, and Microdilutors The MI test may be carried out in containers of any volume but is most conveniently undertaken in disposable plastic plates that contain 96 U-shaped wells, each of which holds approximately 0.2 ml of fluid. These plates are available from several commercial suppliers. Because some of the plates may be coated with substances that are acquired during manufacture and that are inhibitory to mycoplasmas, it may be necessary to wash them in ethanol for a few minutes, rinse in deionized water for 30 minutes, and air dry. A variety of reusable and disposable plastic micropipettes delivering 0.025- to 0.05-ml volumes is available. It is important that the pipettes are clean enough to allow fluid to drop directly from the tip and not accumulate around it as there could be variation in the volume delivered. To ensure cleanliness, reusable pipettes should be rinsed in saline and deionized water, boiled in deionized water for 10 minutes, and air dried. The pipettes should be examined periodically for correct delivery by transferring a drop of 0.9% NaCl solution to an absorbent "GO-NO-GO" test pad. A drop of the correct size should fill completely the ring on the test pad. Stainless-steel diluters that transfer exactly 0.025 ml of fluid are available. They should be examined for correct delivery by dipping in the saline solution and testing as described for the micropipettes.
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Plate Sealing
Plates should be sealed with cellophane tape cut from rolls or with precut pressure-sensitive tape. The latter is available from Cooke Engineering Co. (Alexandria, VA, No. 220-30) or from Falcon Plastics (Oxnard, CA, No. 3044). Filters
Swinnex filters of 220-nm pore diameter are used to filter organism-antibiotic mixtures.
Procedures The procedure for undertaking the MI test for antibiotic activity is a minor modification of that used for measuring antibody activity, the latter having been described in detail before (Senterfit, 1983; Taylor-Robinson, 1983). In brief, a 0.025-ml volume of liquid medium is dropped into each well of the microliter plate. The antibiotic to be tested is added in a 0.025-ml volume to the first well by a micropipetter or microdiluter and diluted subsequently in serial twofold steps. Seven drops (0.175 ml) of the mycoplasma culture, diluted in medium to contain 100-1000 color-changing units in this volume, are then added to each of the wells, except those serving as medium controls. It is helpful to incubate plates at 3TC for 20-30 minutes and then seal rapidly at room temperature or in a more leisurely manner in a 37°C walk-in incubator. This procedure of warming and expanding the medium before sealing helps prevent the tape from loosening on continued incubation of the plate at 37°C. Incubation at 30-32°C is best for tests on spiroplasmas, and room temperature may be used for tests on acholeplasmas and fast-growing mycoplasmas. Indication of Cidal or Lack of Cidal Activity
As indicated earlier, the MIC is the higest dilution of the antibiotic at which the change in color of the medium is inhibited. This should be recorded first when the medium in the wells containing the mycoplasma organisms without any antibiotic has altered in color equivalent to a change of approximately 0.5 of a pH unit. With experience this degree of change may be judged easily by the naked eye. This is a recording of the initial inhibitory concentration (IIC). In most cases, on continued incubation, color changes occur in wells containing successively increasing concentrations of the antibiotic; in other words, there is a "creeping" increase in the MIC value. When the changes cease to occur, the final
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David Taylor-Robinson
inhibitory concentration (FIC) of the antibiotic is recorded. Failure of the antibiotic to kill the organisms is particularly apparent when there is a wide divergence between the IIC and FIC values. However, even when these values are close, it does not necessarily mean that the antibiotic is cidal in its activity. Such activity has to be tested for in one of the following two ways. Cidal Activity Determined by Antibiotic Dilution At the time the IIC is recorded, medium is removed from a well containing organisms only and from wells containing the organism-antibiotic mixtures. The latter media are withdrawn from the wells which contain antibiotic concentrations below, at, and above the IIC value. The contents (0.2 ml) of each of these wells are taken most easily by puncturing the tape with a needle and withdrawing the medium into a 1-ml syringe. This medium is inoculated into 20 ml of fresh medium (a 1:100 dilution) which is then incubated at 37°C or other appropriate temperature to await the development of a color change. If this occurs, confirmation that it is due to the multiplication of mycoplasmas that have not been killed by the antibiotic may be obtained by withdrawing an aliquot and plating it onto the appropriate agar medium for the detection of colonies. If a color change does not occur on continued incubation, it is an indication that the antibiotic in the well from which the organism-antibiotic mixture has been removed had a cidal effect. Cidal Activity Determined by Filtration The same procedure as described earlier is followed initially. Thus, the contents of a well (0.2 ml) are withdrawn with a needle attached to a 1-ml syringe and then 0.8 ml of fresh medium is drawn up into the syringe. The contents of the syringe are passed through a filter of 200-nm pore diameter. The organisms trapped on the filter are "washed" by passing complete medium or broth without the supplements through the filter in, for example, two successive volumes of 5 ml. The filter is then taken from the holder and placed in 20 ml of medium which is incubated at 37°C or other appropriate temperature to seek evidence of a color change. Subsequent procedures are the same as described earlier. In both methods, the lowest concentration of antibiotic at which there is no growth of the mycoplasma is the cidal concentration.
Discussion Antibiotics are effective clinically in treating mycoplasmal infections, despite the fact that generally they are not mycoplasmacidal, because of the cooperation
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of the immune system in eradicating the organisms. When the immune system is defective or suppressed, the fact that most antibiotics do not kill mycoplasmas, at least in concentrations that are achievable in vivo, becomes most apparent. In other words, there is often persistent failure to eradicate the organisms from infected patients with impaired immunity. Such failure is also seen in attempts to eradicate mycoplasmas from cell cultures. The quinolones are regarded as having greater cidal activity than antibiotics that have been used hitherto for the treatment of mycoplasmal infections. This may be noted indirectly by the greater efficiency that such an antibiotic has compared to another in eradicating mycoplasmas from cell cultures and in treating immunodeficient patients. However, direct evidence for the cidal activity of an antibiotic, or the lack of it, may be ascertained only by the methods outlined here. Tests of antibiotic sensitivity based on the inhibition of colony development on agar medium are not suited to determining cidal activity because the antibiotic cannot be separated from the organisms once the test has been set up. In contrast, this may be achieved, as described, when a MI test for antibiotic activity is used. However, the procedures required to assess cidal activity are probably unnecessary when there is rapid "breakthrough" of color changes in a MI test; in other words, when the change from the IIC value to the FIC value occurs rapidly and the latter becomes widely divergent from the former. This has been seen, for example, in tests of rosaramicin (Taylor-Robinson and Furr, 1982) and is in stark contrast to the observations in tests of povidone-iodine, where the closeness of the IIC and FIC values indicated cidal activity (Furr and Taylor-Robinson, 1980). An aspect worth emphasizing is that the cidal activity of an antibiotic should be determined at the time that color changes in the MI test are early in their evolution, i.e., when the IIC value is recorded. To allow incubation of the organism-antibiotic mixtures to continue and then test is unwise because organisms in the mixtures may have become inactivated thermally rather than through the activity of the antibiotic. Furthermore, it should be noted that organisms in control wells containing organisms only are likely to die sooner than those in wells which also contain a mycoplasmastatic antibiotic. The latter, in contrast to a cidal drug, is likely to maintain viability through preventing the rapid multiplication of organisms and the inevitable death that occurs in the decline phase of a growth curve.
References Freundt, E. A. (1983). Culture media for classic mycoplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 127-135. Academic Press, New York. Furr, P. M., and Taylor-Robinson, D. (1980). The killing of Ureaplasma urealyticum and Mycoplasma hominis by povidone-iodine. J. Antimicrob. Chemother. 6, 225-230. Roberts, M. C. (1992). Antibiotic resistance. In "Mycoplasmas: Molecular Biology and Pathogenesis" (J. Maniloff, R. N. McElhaney, L. R. Finch, and J. B. Baseman, eds.), pp. 513-523. Am. Soc. Microbiol., Washington, DC.
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Senterfit, L. B. (1983). Tetrazolium reduction inhibition. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.), Vol. 1, pp. 419-421. Academic Press, New York. Shepard, M. C. (1983). Culture media for ureaplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.), Vol. 1, pp. 137-146. Academic Press, New York. Taylor-Robinson, D. (1967). Mycoplasmas of various hosts and their antibiotic sensitivities. Postgrad. Med. J. 43, 100-104. Taylor-Robinson, D. (1983). Metabolism inhibition tests. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.) Vol. 1, pp. 411-417. Academic Press, New York. Taylor-Robinson, D., and Furr, P. M. (1982). The static effect of rosaramicin on Ureaplasma urealyticum and the development of antibiotic resistance. J. Antimicrob. Chemother. 10, 185191. Whitcomb, R. F. (1983). Culture media for spiroplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.). Vol. 1, pp. 147-158. Academic Press, New York.
SECTION
D
Diagnosis of Specific Diseases
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Dl INTRODUCTORY REMARKS Joseph G. Tully
Diagnostic techniques for mollicutes have literally undergone a major revolution since the earlier published methodology series on these organisms (Razin and Tully, 1983; Tully and Razin, 1983). The changes include a significant expansion in the range of hosts colonized or infected with these organisms, in the number of newly characterized mollicute species, and the application of new molecular techniques for the detection and identification of such organisms. The procedures presented in this section record developments that have occurred in this area. The diagnosis of human Mycoplasma pneumoniae infection has been seriously compromised by the nonspecificity of the conventional complement fixation test and obvious impediments to a rapid diagnosis based on cultural procedures. The feeling still persists that the delay in prompt diagnosis and institution of proper treatment plays a role in the occurrence of some of the serious extrapulmonary infections observed. Chapter D2 and also Chapter B5 in this volume review the current status and evaluation of the newer diagnostic methods for the measurement of a specific antibody in the infection. The discussion also covers the advantages of various antigen detection techniques in providing early diagnosis of infection. The problem in establishing a role for mycoplasmas in human sexually transmitted diseases has always been complicated by the commensal mycoplasma flora in the normal human urogenital tract. Chapter D3 outlines important criteria for delineating etiologic involvement of such organisms and presents clinical and laboratory approaches for a proper diagnosis. The role of mycoplasmas in human immunodeficiency virus (HIV) infections and in development of AIDS (acquired immunodeficiency syndrome) still remains a puzzle. The documentation of specific Mycoplasma species in such hosts and the ability 207 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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of these organisms to alter major host immune responses (see Chapters F5 to Fl 1 in Vol. I), are associations not well understood or explained. In Chapter D5, the authors summarize the current status of the four major Mycoplasma species found in AIDS patients. It is hoped that further application of a combined sensitive and specific detection system, based on DNA amplification techniques and careful epidemiologic studies, will eventually provide an understanding as to the role these organisms have in this human disease. In contrast, careful studies have clearly established an important role for two sexually transmitted mollicutes (Ureaplasma urealyticum and M. hominis) in human neonatal infections. Chapter D4 reviews the occurrence of pneumonia and meningitis, especially as invasive infections in preterm infants, and details appropriate laboratory techniques for proper diagnosis. The widespread occurrence of animal mollicute infections with their important economic impact has continued to stimulate the development of diagnostic techniques that define new mollicute species and promote rapid detection procedures in the hosts. Chapter D6 reviews a variety of culture methods and serologic tests that can be used in diagnoses of bovine mycoplasmas. Improvements in the serologic detection of bovine pleuropneumonia infections are outlined in Chapter B3 of this volume. Bovine semen contamination with mycoplasmas is a particular problem in both localized and geographic dissemination of infection through artificial insemination programs. Suitable diagnostic tests for such contamination are also outlined in Chapter D6. Diagnostic tests for caprine and ovine mycoplasma infections (Chapter D7) have been marked by an expansion in a number of newly characterized mollicute species, especially those inhabiting the caprine ear. Culture methods are usually successful in the diagnosis of colonization or infection in both goats and sheep, with urogenital and mammary gland infections being most commonly observed. In the diagnosis of caprine pleuropneumonia infection, serologic techniques and some antigen detection methods have been refined to detect the newly described etiologic agent F38 (see earlier discussion in Chapter B3, this volume). Diagnostic techniques for swine mycoplasma infections are presented in Chapter D8. Some of the detection methods based on culture procedures are complicated by mixed infections, such as the overgrowth by M. hyorhinis in cultivation attempts for the slower-growing and more fastidious M. hyopneumoniae. Application of a direct immunofluorescence test on impression smears from porcine lung tissue seems to provide a rapid clinical diagnosis of enzootic pneumonia infections due to M. hyopneumoniae. Supporting diagnostic tests based on serologic analysis (see also Chapter B3) and on molecular probes are also presented in this chapter. Diagnostic approaches to mycoplasmas of avian origin (Chapter D9) have also been marked by a significant increase in the number of new mollicute species and in an expansion of involved hosts, especially in birds of prey (raptors). Since birds harbor a considerable flora of commensal mollicutes, diagnostic techniques are more frequently directed to the
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detection of pathogenic species on a flock basis. Emphasis has been directed to more rapid diagnostic procedures through molecular probes, DNA amplification (polymerase chain reaction, PCR) techniques, or enhanced serologic techniques (see Chapter B2) for the major pathogens in the group. The detection of mollicutes in plant and insect infections covers an extensive host range, with relevance to a multitude of different mollicutes, representing as many as five distinct genera in the class Mollicutes. Current diagnostic approaches to Spiroplasma citri infections in plants and insects (Chapter DIO) center around a rapid and specific PCR technique, which is advantageous in detecting plant infections prior to symptom expression. Apparently, molecular techniques have yet to be applied to S. kunkelii infections in com. The principal diagnostic approach in such infections involves either conventional culture or an ELISA sandwich technique, using a polyclonal antiserum. The diagnosis of phytoplasma infections of plants (Chapter D l l ) has been complicated by an enormous complexity in the number and genetic variability of the etiologic agents involved (see Chapters B2 and E6 in Vol. I). Developments from an international collaborative program on the taxonomy and phylogeny of these organisms (Chapter E6, Vol. I) have provided the basis for a more satisfactory and effective identification scheme. One effective early approach to the problem, as reported in the chapter, involves a rapid but nonspecific direct detection of organisms in plant material using a DNA-binding fluorochrome dye. This test still has some practical diagnostic value as a preliminary screening procedure. However, DNA hybridization, including both dot-blot and Southern blot analyses, and PCR amplification techniques can now provide a suitable grouping and differentiation scheme for the approximately 51 currently described, noncultivable, plant-pathogenic phytoplasmas. Detection and identification techniques for various Spiroplasma, and Acholeplasma, Mesoplasma, and Entomoplasma species in insect hosts are described in Chapter D12. Conventional cultural and serologic methods usually suffice for isolation and identification of these mollicutes. However, mixed spiroplasma infections are frequently observed in insect hosts, such as tabanids (horseflies), complicating many conventional serologic procedures (metabolism inhibition and growth inhibition) that require purified antigens for species identification. The chapter presents various advantages of the direct spiroplasma deformation test in detection and separation of such mixed infections.
References Razin, S., and Tully, J. G., eds. (1983). "Methods in Mycoplasmology," Vol. 1. Academic Press, New York. Tully, J. G., and Razin, S., eds. (1983). "Methods in Mycoplasmology," Vol. 2. Academic Press, New York.
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D2 LABORATORY DIAGNOSIS OF Mycoplasma pneumoniae INFECTION R. J. Harris^ J. Williamson^ C. Hahn^ and B. P. Marmion
Introduction and Overview The early expectations for laboratory techniques allowing the rapid diagnosis and immediate clinical management of atypical pneumonia caused by Eaton agent (Mycoplasma pneumoniae), which were raised by the isolation of the causative organism (Eaton et aL, 1944), its identification as a mycoplasma, and its growth in cell-free media (Marmion and Goodbum, 1961; Chanock et aL, 1962), and the subsequent development of various antibody assays, are only now nearing reality. However, there is no single test or array of tests which will provide a rapid, economical diagnosis which can direct clinical chemotherapeutic decisions in a reasonable time frame (:^24 hours) and also detect a realistic percentage of infections (>:90%), although some tests are approaching that ideal. One established test is the antigen capture or antigen-enzyme immunoassay (Ag-EIA) of Kok et al. (1988). Simplified, quantitative polymerase chain reaction (PCR) DNA amplification, now under development in a number of centers, will no doubt provide another rapid test. The development of effective tests has been hampered by two major constraints. First, and principally, confirmation by culture of the organism is lengthy and difficult. The difficulty in culturing the organism presumably arises from its minimal absorptive and synthetic capability. Few other major human pathogens offer similar difficulties. Even with improved diphasic medium (Kenny et al., 211 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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1990), the growth, subculture onto solid medium, and identification of colonies may take 7 to 42 days. Moreover, even when completed evidence shows that culture only detects a proportion (68%) of those infected as judged by serodiagnosis. Second, and conversely, serodiagnosis by the commonly used complement fixation test is also slow. The conventional criterium for identification of a current infection is a fourfold rise in specific antibody (total or IgM generally) and requires collection of paired sera over an interval of 5-8 days. Additionally, serodiagnosis falls short of detecting all culture-positive patients. Thus both culture and serodiagnosis are too slow to direct early clinical intervention. Despite the limitations, the routine laboratory diagnosis of M. pneumoniae infection often relies on serodiagnosis alone, principally by complement fixation or, more recently, on EI A techniques or agglutination of antigen-coated particles. In many clinical settings a tentative diagnosis of M. pneumoniae infection is based on a number of clinical and epidemiological factors (Clyde, 1993). These selective factors are at present the mainstay of diagnosis because they allow rapid chemotherapeutic intervention. Strategies for laboratory diagnosis which have been developed (but not necessarily clinically assessed or widely available commercially) are summarized in Fig. 1 and Table I. Three categories of tests have been devised. These comprise first, direct detection of the organism by culture; second, detection of cellular components (DNA, rRNA, antigens); and third, quantification of the nonspecific and specific immunological responses of infected individuals. Direct methods comprise culture of the organism via liquid culture, agar, diphasic, and cell sheet. Methods for detection of cellular components involve detection of the antigens of the organism (total proteins, PI adhesin, or glycolipids), detection of specific DNA sequences, and detection of specific ribosomal RNA sequences. Assays of the immunological response of infected individuals include those for nonspecific cold hemagglutinins; M. pneumoniae-specific IgM, IgG, and IgA; and a general (nonspecific) T-cell activation marker, i.e., raised serum adenine deaminase activity. The three categories of tests are presented in detail in the following discussion.
Detection of Cellular Components of M. pneumoniae Detection of M. pneumoniae Antigens by El A Kok et al. (1988) have developed a simple and effective antigen capture method (Ag-EIA) for detecting M. pneumoniae antigens (proteins) in respiratory exudates. The antigen capture assay protocol involves first coating of microtiter wells
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D2 Laboratory Diagnosis of M. pneumoniae
rRNA
M. pneumoniae
t
Culture 1 Liquid Culture 2 Agar 3 Diphasic 4 Cell Sheet
In-Solution Hybridisation Assay (Gen-Probe)
P1 Adhesin
Immune Response
IgM igG igA IgE
Antik)ody Assays 1 CF 2 IHA(t) 3 IHA (M) 4 IgM-EiA 5 IgA-EIA 6 IgE-EIA
DNA
Antigen Assays 1 Immunofluorescence ^ 2 Antigen-EIA (Ag-capture) 3 P1 specific EIA 4 T cell response (raised adenine deaminase) DNA Assays 1 Dot blot hybridisation 2 PCR - with DNA detected by: (a) Agarose gel electrophoresis (b) Dot blot hybridisation (c) Solid phase capture (d) Solid phase PCR
Fig. 1. Listings of diagnostic targets of M. pneumoniae and corresponding assays for detection of infections mediated by the organism.
with "capture" rabbit antiserum to M. pneumoniae. A point of central importance is the adsorption of the rabbit antiserum with human fetal lung. Specimens are diluted in skim milk (or casein)-phosphate-buffered saline (PBS)-Tween, sonicated briefly, and then added to the coated wells. The detector layer consists of guinea pig antisera to M. pneumoniae followed by the rabbit anti-guinea pig sera coupled to horseradish peroxidase (indirect kg capture). Substitution of a PI adhesin-specific, monoclonal antibody either for "capture," or in the detection system, was less effective. Similarly, the use of a high-titer polyclonal, Plspecific, rabbit antiserum did not improve sensitivity. The latter was raised by transfection of RK13 rabbit cells with a "universal" coding equivalent of the PI adhesin gene (i.e., UGA terminator codons removed by in vitro mutagenesis;
TABLE I PRINCIPAL FEATURES OF TESTS FOR DETECTION OF M. pneumoniae INFECT~ON Indicator of M. pneumoniae infection: Detection of Live organisms
Total antigenic mosaic of M. pneumoniae: live or dead organisms or fragments
PI surface adhesion DNA
Ribosomal RNA (16s)
Test procedure Liquid culture Agar culture Disphasic medium Celt sheet culture Direct or indirect immunofluorescence
Type of information generated
Test time 10-20 days
Retrospective
Percentage (max) of infectionsdetecteda
Comments
-40 68 (ref. 3)
5 days -2 hours
Current
86 (ref. 1)
Interpretation of immunofluorescence patterns difficult
Direct or indirect Ag-EIA
-24 hours
Current
59 (ref. 5)
Ag-EIA Dot-blot hybridization PCR-amplified DNA detected by (a) Agarose gel electrophoresis (b) Dot-blot hybridization (c) Solid phase capture (d) Solid-phase PCR In solution hybridization with '251 probe
-4 hours 3.5 hours
Current Current
-
Collection of NPAb unpleasant for adults. Sensitivity -lo4 CFUIml Low sensitivity with NPA Sentitivity 105 CFU
3.5 hours
Current
5 hours 4 hours 3.5 hours 3 hours
Current Current -
Current
38 (ref. 4)
-
Very laborious -40 (ref. 10) 83 (ref. 7)
Stored specimens used Sensitivity poor at present No longer available from Gen-Probe
Immunological indicators Cold agglutinins
IgM antibody to M . pneumoniae
IgG to M. pneumoniae
IgA to M. pneumoniae
Total antibody to M. pneumoniae Adenine deaminase in blood (nonspecific T-cell activation marker)
Agglutination of patients or group 0 erythrocytes (a) Complement fixation (CF)
3-6 hours or overnight Overnight
Current, even bedside
30-50 (ref. 2)
Early serological indicator of infection, but not specific
Current (presumptive) for one serum, retrospective for paired sera
Less sensitive than (b) -40 (ref. 9) current
-
(b) EIA: p. chain capture or antigen bound to solid phase
-
(c) Modified indirect hemagglutination: p. chain capture IHA(M) EIA with antigen bound to beads or cups in plates. Agglutination of latex or gelatin beads coated with antigen EIA with antigen bound to beads or cups in plates. Agglutination of latex or gelatin beads coated with antigen Indirect hemagglutination IHA(t)
-
-
-90 (ref. 6)
24 hours
Current (presumptive) for one serum, retrospective for paired sera
-
Demonstration of an increase in titer is required
24 hours
Current (presumptive) for one serum, retrospective for paired sera
-
Demonstration of an increase in titer is required
2-3 hours
Current (presumptive) for one serum, retrospective for paired sera Current
? (ref. 6)
More sensitive than CF
? (ref. 8)
Most bacterial pneumonias gave negative values
Adenine deaminase enzymatic assay (of serum)
2 hours
With assays (b) and (c), a moderate or high level or titer of IgM antibody is highly suggestive of recent or current infection: increasing levels are definitive for current infection.
UI
a 100% of infections usually ascertained by serological means. Key to references: (1) Hirai et al. (1991); (2) Jacobs (1993); (3) Kenny et al. (1990); (4) Kleemola et al. (1993); (5) Kok et al. (1988); (6) Kok et al. (1989); ( 7 ) Luneberg et al. (1993); (8) Suga et al. (1989); (9) Samra and Gadba (1993); and (10) Williamson et al. (1992) bNasopharyngeal aspirate.
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Professor P.-C. Hu, University of North Carolina) under the control of a zincinducible metallothionein promoter. Zinc induction of the PI gene led to the production of PI protein which was then used (unpurified) to raise high titer Plspecific antibodies in rabbits (Williamson et al, 1994). Attempts to express the PI gene in Escherichia coli have been unsuccessful. Of a wide range of mycoplasmas and bacteria (many of which can be found in the respiratory tract), only M. genitalium reacted to a minor extent in the AgEIA as devised by Kok et al. (1988). The assay detected 10"^ colony-forming units (CFU)/ml in specimens artificially "laced" with known numbers of M. pneumoniae. The Ag-EIA detects 59% of serologically proven cases; about the same rate as improved culture reported by Kenny et al. (1990). The 100% or reference "gold" standard was provided by serodiagnosis, either a high, unchanging CF antibody titer with IgM detected by hemagglutination-IgM capture (see later) or a fourfold or greater rise in specific antibody titer. Antigen capture-EIA is much quicker (18 hours) than culture and provides an initial diagnosis for clinical management, with later consolidation of the diagnosis by serological examination. A similar Ag-EIA has been developed by Kleemola et al. (1993) and is marketed as Enzygost by Behring. Since 1988, in the virus diagnostic laboratory of the Institute of Medical and Veterinary Science, Adelaide, the Ag-EIA for M. pneumoniae has been included with other Ag-EIA for direct diagnosis of infection due to respiratory viruses. This has been highly effective in detecting M. pneumoniae infections in the late winter/spring (southern hemisphere) of 1987, 1988, 1989, and 1990, but with only sporadic cases in 1991. Detection of M. pneumoniae Antigens by Immunofluorescence Understandably, modem microbiological laboratories are avoiding the more labor-intensive (and therefore expensive), subjective visual immunofluorescencebased diagnostic methods for antigen detection and prefer semiautomated test systems which are principally EIA in nature. Nevertheless, an indirect IF test, based on previous observations by Hers and Masurel (1967), has been successfully reexplored by Hirai etai (1991). Smears made from throat swabs were examined with a carefully absorbed polyclonal rabbit antiserum against M. pneumoniae and samples from 42/49 (86%) patients with serological evidence of current infection were positive. Detection of M. pneumoniae by Probing for Specific Nucleotide Sequences Cloned genomic fragments of M. pneumoniae have been used by us and others (Hyman et al., 1987) in dot-blot hybridization-based assays for detection of
D2 Laboratory Diagnosis of M. pneumoniae
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M. pneumoniae. In our laboratory, two M. pneumoniae-spQcific probes, labeled with 32P, comprising ~1000-bp double-stranded (denatured) molecule and a single-stranded 500 nucleotide Ml3 probe were prepared and tested (R. Harris et al., unpublished). However, their limit of detection, 10^ CFU, of M. pneumoniae was considered too insensitive for further development (see Chapters Al and A3, this volume; and review Razin, 1994). Attention was therefore turned to the M. pneumoniae rapid diagnostic system (Gen-Probe, San Diego, CA). This assay involves an "in solution" hybridization of a i25i.iabeled probe against a specific ribosomal RNA (rRNA) sequence of the mycoplasma. The assay was considerably more sensitive with simulated samples than was Ag-EIA or hybridization with the DNA probes, presumably reflecting the larger number of target rRNA molecules versus genomes per cell; —10^ CFU of M. pneumoniae was detected. However, rather surprisingly with clinical specimens, the rapid diagnostic system detected only about one-fourth of the specimens detected by Ag-EIA. This insensitivity was speculatively attributed to selective degradation (or rapid selective clearance) of the target rRNA in respiratory secretions during natural infections, once the membrane of the organism had been breached by immune reactions or chemotherapy. Proteins and glycolipids detectable by Ag-EIA were speculated to be cleared from the respiratory tract more slowly (Harris et al., 1988). In view of these findings, attention was turned to PCR amplification of sequences within two genes of M. pneumoniae as a method of direct detection of the organism. Primers and probes (described in detail in Williamson et al., 1992) were designed to detect a section of the PI or cytadhesin gene with the product identified by dot-blot hybridization (DBH) following PCR amplification (PlPCR-DBH assay). Similarly, sequences within the 16S rRNA gene were assayed (16S rDNA-PCR-DBH assay). The latter assay was established to exclude certain antigen-positive, PCR-negative results that may arise from a possible strainto-strain variation in the PI gene; it is presumed that the rRNA gene sequence would be highly conserved between strains of M. pneumoniae. The PCR product from each assay was detected and quantified by DBH with a synthetic ^^p. labeled hairpin probe. The bound counts in the blots were quantified by scintillation counting, and a sample ratio was derived: sample ratio = (sample counts per minute - background counts per minute)/background counts per minute (Williamson et al., 1992). The sensitivity was excellent; 50 CFU of M. pneumoniaelmX were detected: 200-fold less than could be detected by Ag-EIA. Tests with a panel of organisms including M. genitalium showed no crossreactions. A quantification curve was constructed from the Pl-PCR-DBH sample ratios obtained from dilutions of a laboratory culture of M. pneumoniae, which, in turn, allowed an approximation of CFU of M. pneumoniaelml in the range of 50-»10^ CFU/ml for any clinical sample. Certain samples were also tested with a PCR-DBH assay for M. genitalium, constructed along the same lines to ex-
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R. J. Harris ef a/.
elude the possibility that Ag-EIA-positive, PCR-DBH-negative samples were the result of the presence of M. genitalium, which cross-reacts antigenically with M. pneumoniae. The antigen-positive/PCR negative samples proved negative for M. genitalium. However, the latter organism was detected in samples from two other subjects negative for M. pneumoniae. While this study was in progress, other investigators developed PCR-based assays for M. pneumoniae mostly using simulated positive samples. Those working with material from naturally infected subjects include de Barbeyrac et at. (1993) and Luneberg et al. (1993). The latter detected, via PCR, 83% of serologically and/or culture proven infections (see Chapter A6, this volume; and Razin, 1994).
Isolation of M. pneumoniae by Culture Considerable advances have been made in improving both the percentage of isolations (i.e., relative to the "gold" standard of serologically proven infections) and in reducing the time in culture to obtain a positive result. Improvements have included development or application of special liquid media (SP4; Tully et al., 1979; Chapter A2, Vol. I) and diphasic (liquid-solid) media; the latter improving isolation by 26% to an impressive 68% of those with serological evidence of infection (Kenny et al., 1990). Also, culture on live but cycloheximide-inhibited feeder cell sheets (Marmion et al., 1993) reduced culture time to as little as 5 days. Detection of M. pneumoniae in the cell sheet system is by immunofluorescence, or preferably Ag-EIA. Direct comparisons of culture in the cell sheet system and in diphasic media with SP4 as the liquid phase were carried out with homogenized lung samples from artifically infected guinea pigs. These comparisons consistently demonstrated that the cell sheet culture was slightly more sensitive and certainly more rapid. However, the same study showed that Pl-PCR-DBH assay was considerably more sensitive and exposed the poor plating efficiency of both culture systems (Marmion et al., 1993).
Detection of M. pneumoniae Infections by Measurement of the Immunological Response of Individuals with Respiratory Infection A number of assays have been used for detection and quantification of M. pneumoniae antibodies, including the older ones based on complement fixation (CF), metabolic inhibition (e.g., tetrazolium reduction-inhibition technique), mycoplasmacidal test, radioimmunoprecipitation, radioimmunoassay, and hemagglutination (Taylor-Robinson et al., 1966; Busolo and Meloni, 1983; Hu et al., 1993) and the newer techniques based on EI A (Jacobs, 1993). Those relying on
D2 Laboratory Diagnosis of M. pneumoniae
219
culture are too cumbersome and slow for modem diagnostic services. However, CF (which predominantly detects IgM) and EIA are widely used and particle agglutination assays have become available. Measurement of IgM and Other Immunoglobulin Class Responses by Modified Indirect Hemagglutination Assay (IHA) (Kok et aL, 1989) Sheep erythrocytes are glutaraldehyde fixed, tanned, and coated with sonicated M. pneumoniae. Sensitized erythrocytes can be stored up to 2 months at 4°C without loss of activity. The assay is based on the antibody capture principle. Microtiter plates are coated with antisera to human |JL, a, or 7 chains, dilutions of the patients' sera are added and incubated, and the wells are subsequently washed. When antigen-coated erythrocytes are added, the settling pattern in those wells in which immunoglobulins to M. pneumoniae have been captured leads to the cells forming a shield rather than a central button in the cup or well. The |x chain capture version of the assay [IHA(M)] has been shown to be consistently more sensitive than the CF test. For example, Kok et al. (1989) found that with the CF assay as the standard, IHA(M) detected 89% of the patients who had CF antibody, whereas in reverse, with IHA(M) as the standard, the sensitivity of the CF test was 64%. Although the results of the IHA(M) assay correlated well with those of a locally developed solid-phase indirect IgM-EIA in formal comparisons, investigations of respiratory infection in the Newcastle survey (see later) showed that IgM-EIA tests were positive for only 2 of 24 subjects who were seropositive by IHA(t) (for total specific antibody) and also by IHA(M). Finally, the combined use of IHA(t)—in essence the test as devised by Dowdle and Robinson (1964) which does not utilize antibody capture—for measurement of the total antibody along with the 7 and a chain capture variants would allow additionally the detection of IgG or IgA responses in the absence of IgM responses in persons who are experiencing reinfection with M. pneumoniae (see Hu et al., 1983; Jacobs, 1993). However, it is possible that the use of antibody capture for the measurement of responses of these immunoglobulin classes might be more sensitive than IHA(t); this requires investigation. Other workers have developed EIA for M. pneumoniae-spQcific IgM (Samra and Gadba, 1993; Wreghitt and Sillis, 1985). These also offer greater sensitivity than the CF test. Care is required to eliminate false IgM positives arising from the presence of rheumatoid factor complexes with M. pneumoniae-specific IgG. Suspect sera can be adsorbed with an IgG-specific latex bead adsorbent. Bead-Based Agglutination Tests for Detection of M. pneumoniae-Specific IgG A high density particle agglutination test for M. pneumoniae IgG has been developed. The test is very rapid (30 minutes) and is more sensitive than the CF
220
R. J. Harris et al.
test (Shitara et al., 1990). However, a latex agglutination test was assessed by Karppelin et al. (1993) and found to be insensitive.
Cross-Reaction of M. genitalium in M. pneumoniae Tests M. genitalium is occasionally found in the respiratory system and shares antigens with M. pneumoniae (Baseman et al., 1988; Lind et al., 1984). M. pneumoniae antibodies have been shown in laboratory tests to cross-react with M. genitalium. Consequently, strictly speaking, a definitive diagnosis cannot be made with most available tests based on the detection of M. pneumoniae antigens or antibodies. However, PCR-based tests are specific. This is expected in view of the small degree of total genomic sequence homology between the two organisms.
Concluding Remarks One of the problems with the laboratory diagnosis of M. pneumoniae infection lies with the well-known insidious nature of disease onset. The patient may not seek medical attention for days or weeks until increasing malaise and respiratory distress force a visit. By that time the initial phase of the respiratory infection may have passed when organisms might have been more easily cultivated or demonstrated. Also, the serological response may have reached a plateau, excluding the possibility of meeting the conventional criterion for a current infection that a fourfold or greater increase in antibody titer must be demonstrated. Several rapid tests are available to assist clinical management: first, antigen detection in respiratory secretions by enzyme immunoassay (Kok et al, 1988; Kleemola et al., 1993) and second, a number of alternative techniques for detection of M. pneumoniae-spQcific total or IgM or other immunoglobulins, early in infection from a single serum sample. Presumptive evidence of infection is indicated by a significant titer, for example, a titer >:32 via CF (Jacobs, 1993) or via EI A or |x chain-specific indirect hemagglutination (Kok et al., 1989). Similarly, detection of early M. pneumoniae-spQcific IgA levels, particularly in cases of reinfection, may be useful. In our experience one of the more convenient and rapid diagnostic procedures is the Ag-EIA of Kok et al. (1988): this detects the "complete" antigenic mosaic of the organism. In its present usage in Adelaide it provides routine detection of the M. pneumoniae antigen in secretions of patients with severe respiratory infections which might be due to M. pneumoniae. Results are timely (18 hours) for directing appropriate chemotherapy. Specificity is excellent, and approximately 59% of serologically proven cases are detected.
D2 Laboratory Diagnosis of M. pneumoniae
221
Surprisingly, PCR-based tests, although more rapid, do not always detect a greater percentage of cases (Williamson et ai, 1992). However, Luneberg et aL (1993) detected 83% of culture and/or serologically proven infections using PCR coupled with anchored liquid-phase hybridization detection of the amplified DNA product. In view of the problems associated with PCR—principally the ease of occurrence in false positives due to contamination of laboratories with amplicons arising from previous amplifications—it may be premature to embark on routine screening with this method. Instead, PCR may be used for the more serious systemic infections (e.g., of the central nervous system) or for infections in immunocompromised individuals with respiratory infections. As stated, serological examination of patients is the predominant method for diagnosis of M. pneumoniae infection. Detection of diagnostically significant levels of total antibody to M. pneumoniae and specific IgM and IgG (and perhaps IgA) from a single blood sample taken from patients shortly after presentations also provides timely presumptive information for clinical decisions on the form of appropriate chemotherapy. According to Granstrom et al. (1994), M. pneumoniae-^^tciiic IgA (via EIA) develops more often and more rapidly than IgM. Additionally, elevated IgM is less often seen in infections in older age groups. For IgM, approximately 40% of infected individuals were positive in the first serum sample (Samra and Gadba, 1993). This percentage is higher if all three antibody classes are determined (Granstrom et al., 1994). However, sensitive M. pneumoniae-specific IgM capture methods are required (e.g., EIA); the conventional complement fixation test is less sensitive. There are therefore a range of diagnostic methods offering improved speed and sensitivity which potentially allow timely chemotherapeutic intervention in M. pneumoniae infections.
References Baseman, J. B., Dallo, S. F., Tully, J. G., and Rose, D. L. (1988). Isolation and characterization of Mycoplasma genitalium strains from the human respiratory tract. J. Clin. Microbiol. 26, 22662269. Busolo, F., and Meloni, G. A. (1983). Serodiagnosis of M. pneumoniae infections by enzyme-linked immunosorbent assay (ELISA). Yale J. Biol. Med. 56, 517-521. Chanock, R. M., Hayflick, L., and Barile, M. F. (1962). Growth on artificial medium of an agent associated with atypical pneumonia and its identification as a PPLO. Proc. Natl. Acad. Sci. U.S.A. 48, 41-49. Clyde, W. A., Jr. (1993). Clinical overview of typical Mycoplasma pneumoniae infections. Clin. Infect. Dis. 17(Suppl. 1), S32-S36. de Barbeyrac, B., Bemet-Poggi, C , Febrer, F., Renaudin, H., Dupon, M., and Bebear, C. (1993). Detection of Mycoplasma pneumoniae and Mycoplasma genitalium in clinical samples by polymerase chain reaction. Clin. Infect. Dis. 17(Suppl. 1), S83-S89. Dowdle, W. R., and Robinson, R. Q. (1964). An indirect haemagglutination test for diagnosis of Mycoplasma pneumoniae infections. Proc. Soc. Exp. Biol. Med. 116, 947-950.
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Eaton, M. D., Meiklejohn, G., and van Herick, W. (1944). Studies on the etiology of primary atypical pneumonia: A filterable agent transmissible to cotton rats, hamsters and chick embryos. J. Exp. Med. 79, 649-668. Femald, G. W., and Clyde, W. A., Jr. (1989). Epidemic pneumonia in university students. J. Adolesc. Health Care 10, 520-526. Granstrom, M., Holme, T., Sjogren, A. M., Ortqvist, A., and Kalin, M. (1994). The role of IgA determination by ELISA in the early serodiagnosis of Mycoplasma pneumoniae infection, in relation to IgG and |x-capture IgM methods. J. Med. Microbiol 40, 288-292. Harris, R., Marmion, B. P., Varkanis, G., Kok, T., Lunn, B., and Martin, J. (1988). Laboratory diagnosis of Mycoplasma pneumoniae infection. 2. Comparison of methods for the direct detection of specific antigen or nucleic acid sequences in respiratory exudates. Epidemiol. Infect. 101, 685-694. Hers, J. F. P., and Masurel, N. (1967). Infection with Mycoplasma pneumoniae in civilians in the Netheriands. Ann. N. Y. Acad. Sci. 143, 447-460. Hirai, Y., Shiode, J., Masayoshi, T., and Kanemasa, Y. (1991). Application of an indirect immunofluorescence test for detection of Mycoplasma pneumoniae in respiratory exudates. J. Clin. Microbiol. 29, 2007-2012. Hu, P . - C , Powell, D. A., Albright, F., Gardner, D. E., Collier, A. M., and Clyde, W. A., Jr. (1983). A solid-phase radioimmunoassay for detection of antibodies against Mycoplasma pneumoniae. J. Clin. Lab. Immunol. 11, 209-213. Hyman, H. C , Yogev, D., and Razin, S. (1987). DNA probes for detection and identification of Mycoplasma pneumoniae and Mycoplasma genitalium. J. Clin. Microbiol. 25, 726-728. Jacobs, E. (1993). Serological diagnosis of Mycoplasma pneumoniae infections: A critical review of current procedures. Clin. Infect. Dis. 17(Suppl. 1), S79-S82. Karppelin, M., Hakkarainen, K., Kleemola, M., and Miettinen, A. (1993). Comparison of three serological methods of diagnosing Mycoplasma pneumoniae infection. J. Clin. Pathol. 46, 1120-1123. Kenny, G. E., Kaiser, G. G., Cooney, M. K., and Foy, H. M. (1990). Diagnosis of Mycoplasma pneumoniae pneumonia: Sensitivities and specificities of serology with lipid antigen and isolation of the organism on soy peptone medium for identification of infections. J. Clin. Microbiol. 28, 2087-2093. Kleemola, M. Raty, R. Karjalainen, J., Schuy, W., Gerstenecker, B., and Jacobs, E. (1993). Evaluation of an antigen-capture enzyme immunoassay for rapid diagnosis of Mycoplasma pneumoniae infection. Eur. J. Clin. Microbiol. Infect. Dis. 12, 872-875. Kok, T., Varkanis, G., and Marmion, B. P. (1988). Laboratory diagnosis of Mycoplasma pneumoniae infection. 1. Direct detection of antigen in respiratory exudates by enzyme immunoassay. Epidemiol. Infect. 101, 669-684. Kok, T., Marmion, B. P., Varkanis, G., Worswick, D. A., and Martin, J. (1989). Laboratory diagnosis of Mycoplasma pneumoniae infection. 3. Detection of IgM antibodies to M. pneumoniae by a modified indirect haemagglutination test. Epidemiol. Infect. 103, 613-623. Lind, K., Lindhart, B. O., Schiitten, H. J., Blom, J., and Christiansen, C. (1984). Serological crossreactions between Mycoplasma genitalium and Mycoplasma pneumoniae. J. Clin. Microbiol. 20, 1036-1043. Luneburg, E., Jensen, J. S., and Frosch, M. (1993). Detection of Mycoplasma pneumoniae by polymerase chain reaction and nonradioactive hybridization in microtitre plates. J. Clin. Microbiol. 31, 1088-1094. Marmion, B. P., and Goodbum, G. M. (1961). Effect of an organic gold salt on Eaton's primary atypical pneumonia agent and other observations. Nature (London) 189, 247-248. Marmion, B. P., Williamson, J., Worswick, D. A., Kok, T., and Harris, R. J. (1993). Experience with newer techniques for the laboratory detection of Mycoplasma pneumoniae infection: Adelaide, 1978-1992. Clin. Infect. Dis. 17(Suppl. 1), S90-S99.
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Razin, S. (1994). DNA probes and PCR in diagnosis of mycoplasma infections, Mol. Cell. Probes 8, 497-511. Samra, Z., and Gadba, R. (1993). Diagnosis of Mycoplasma pneumoniae infection by specific IgM antibodies using a new capture-enzyme-immunoassay. Eur. J. Epidemiol. 9, 97-99. Shitara, M., Ito, K., Yoshimoto, K., Umezu, S., Kobayashi, S., Itagaki, T., Shitara, M., Kinoshita, T., Nakahara, T., and Hayashi, Y. (1990). Comparative studies of serological test for Mycoplasma pneumoniae on the 73 cases of lower respiratory infection disease. Jpn. J. Clin. Pathol. 38, 683-687. Suga, M., Nishikawa, H., Ando, M., Tanaka, F., Akaike, T., Sakata, T., Kawano, O., Ito, K., Nakashima, H., and Araki, S. (1989). Usefulness of serum adenine deaminase activity in the early diagnosis of Mycoplasma pneumoniae. Nippon Kyobu Shikkan Gakki Zasshi 27, 461466. Taylor-Robinson, D., Sobeslavsky, O., Jensen, K. E., Senterfit, L. B., and Chanock, R. M. (1966). Serologic response to Mycoplasma pneumoniae infection. 1. Evaluation of immunofluorescence, complement fixation, indirect haemagglutination, and tetrazolium reduction inhibition tests for the diagnosis of infection. Am. J. Epidemiol. 83, 287-298. Tully, J. G., Rose, D. L., Whitcomb, R. F., and Wenzel, R. P. (1979) Enhanced isolation of Mycoplasma pneumoniae from throat washings with a newly modified culture medium. J. Infect. Dis. 139, 478-482. Williamson, J., Marmion, B. P., Worswick, D. A., Kok, T., Tannock, G., Herd, R., and Harris, R. J. (1992). Laboratory diagnosis of Mycoplasma pneumoniae infection. 4. Antigen capture and PCR-gene amplification for detection of the mycoplasma: Problems of clinical correlation, Epidemiol. Infect. 109, 519-537. Williamson, J., Marmion, B. P., Kok, T., Hu, P. C , and Harris, R. J. (1994). Development of a Mycoplasma pneumoniae antigen capture assay utilizing the PI adhesin gene cloned in a mammaUan cell line lOM Lett. 3, 512-513. Wreghitt, T. G., and Sillis, M. (1985). A |x-capture ELISA for detQciing Mycoplasma pneumoniae IgM: Comparison with indirect immunofluorescence and indirect ELISA. J. Hyg. 94, 217227.
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D3 DIAGNOSIS OF SEXUALLY TRANSMITTED DISEASES David Taylor-Robinson
Introduction This chapter is concerned first with the steps required to determine whether a member of the class Mollicutes is a cause of a sexually transmitted disease. In this regard, the observations are not confined to human disease since the principles are applicable to disease in other species. Second, sexually transmitted human diseases for which a mycoplasmal involvement has been reasonably assured are mentioned, together with other diseases that are not considered to be sexually transmitted, but which are caused by mycoplasmas usually residing in the urogenital tract. This is followed by a more detailed account of the techniques required to determine whether a mycoplasmal infection has occurred and, therefore, whether it might be involved etiologically in the human disease. Criteria to Be Fulfilled for Etiological Involvement The following are guidelines that need to be adhered to in deciding whether a mycoplasma is one of the causes of a sexually transmitted disease. It should be determined whether (i) the isolation rate for patients with the disease is significantly greater than that for subjects without the disease; (ii) more organisms of the particular mycoplasmal species are recovered from patients with the disease than from subjects without the disease; (iii) an antibody response to the mycoplasma occurs in patients with the disease and that it occurs significantly more 225 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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often in them than it does in subjects without the disease; (iv) there is a clinical response to an antibiotic to which the mycoplasma is susceptible in vitro and the response is accompanied by elimination of the mycoplasma; (v) an antibiotic which inhibits the mycoplasma but not other putative causal agents (a "differential" antibiotic) produces a beneficial clinical effect; (vi) the mycoplasma, when introduced into an animal species, produces a disease that is similar to that in the human, the induced disease is associated with an antibody response to the mycoplasma, and is susceptible to antibiotic therapy; (vii) the mycoplasma, when given experimentally to human volunteers, produces a disease that is similar to that occurring naturally, the induced disease is associated with an antibody response to the mycoplasma, and is susceptible to antibiotic therapy; and (viii) a specific mycoplasmal antibody that is induced naturally or through immunization protects against development of the disease. Sexually Transmitted Diseases Known to Be Caused by Mycoplasmas
Many conditions have been associated with infection by mycoplasmas, but few can be considered to have a mycoplasmal cause. These diseases in men and women and the mycoplasmas attributed as a cause are (i) nongonococcal urethritis in men, Ureaplasma urealyticum and Mycoplasma genitaliuin\ (ii) acute epididymitis, U. urealyticum', and (iii) pelvic inflammatory disease in women, M. hominis. Other Diseases Caused by Sexually Transmitted Mycoplasmas
There are several conditions that are not in the usual sense considered to be sexually transmitted but are caused sometimes by mycoplasmas which by virtue of their dominant urogenital tract colonization are sexually transmitted. These diseases and the mycoplasmas involved are (i) pyelonephritis, M. hominis; (ii) postpartum and postabortion fever, M. hominis and U. urealyticum; (iii) pneumonia, chronic lung disease, and meningitis in very low birth weight infants, U. urealyticum; and (iv) arthritis in hypogammaglobulinemic patients and in those on immunosuppressive regimes, U, urealyticum and M. hominis.
Procedures The technical procedures required to determine whether a mycoplasmal infection has occurred have been detailed elsewhere (Taylor-Robinson, 1983a, 1989) and are outlined briefly, with some additional information. The procedures are those concerned with detection of the organisms, namely culture or other means,
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and the measurement of a specific antibody response to them. Considerable care is required in the collection and transport of specimens to the laboratory, particularly if culture is to be attempted. Specimen Collection and Pretest Handling The detection of mycoplasmas and ureaplasmas in the male urogenital tract may be accomplished by collecting and testing an urethral swab specimen and/or a first-catch urine specimen. However, the latter is likely to prove at least 10-fold less sensitive unless the specimen is centrifuged and the cellular deposit is tested. High vaginal and endocervical swab specimens are likely to be more satisfactory than urethral swab or urine specimens for detecting the organisms in women. They should be collected without contact with antiseptics, analgesics, or lubricants that are often used in obstetrical and gynecological practice, some of which may kill mycoplasmas. Swabs from whatever site should be expressed immediately in mycoplasmal or other transport medium, not broken off into the medium, and should never be allowed to dry. Having medium close to the patient is a sound policy. Samples other than swabs should also be inoculated into medium soon after collection. In the case of blood, for example, it is important to do this as the ability to recover genital mycoplasmas declines appreciably over 1 hour; 2 ml of heparinized blood and 18 ml of medium are appropriate volumes in a Universal bottle. If maintenance of viability is important, media used for transporation may be (i) growth medium {vide infra), with or without the substrates metabolized by mycoplasmas and ureaplasmas; (ii) nutrient broth enriched with, for example, 10 to 20% horse serum; or (iii) sucrose-phosphate transport medium (designated 2SP) containing 10% heat-inactivated (56°C for 30 minutes) fetal calf serum, without antibiotics. The latter is used primarily for the transport of chlamydiae but mycoplasmas may also be recovered from the medium (Smith et al., 1977), obviating the need to insert two swabs into the urethra. Transport medium is also available commercially (Kibsey et al., 1991). If viability is unimportant, for example when specimens are required for antigen or DNA detection, then they may be taken and transported in phosphate-buffered saline (pH 7.2). Lack of transport medium requires that a swab or tissue specimen is taken to the laboratory as rapidly as possible; the same applies to a sample of urine. If transportation takes several hours, the specimen should be kept at 4°C, although this may have the undesirable effect of precipitating phosphates from urine. Once a mycoplasmal medium has been inoculated, there is no great urgency for transportation to the laboratory. Nevertheless, this should be done as soon as possible, preferably within 24 hours, and the medium containing the organisms should be kept at 4°C in the meantime. If transportation cannot be undertaken within a few days (5 at a maximum), the medium containing the specimen should be frozen to
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-70°C or placed in liquid nitrogen and transported in the frozen state. Concern over speed of transportation is less if the specimens are to be examined by noncultural methods. Specimens that are received frozen may be kept indefinitely at -70°C (Furr and Taylor-Robinson, 1990) or in liquid nitrogen and examined after rapid thawing in a 37°C water bath. Prompt examination of specimens received unfrozen is preferable because some loss of viability is inevitable through the freezing process. It is feasible but undesirable to keep specimens at 4°C for a week or longer. If it is clear that testing is not possible within this time, they should be stored as indicated earlier, but not at — 20°C. Isolation Procedures GROWTH MEDIA
The isolation of many mycoplasmas has depended on a medium comprising beef heart infusion broth, available commercially as PPLO broth, supplemented with 10% (v/v) fresh yeast extract (25%, w/v) and 20% (v/v) horse serum (Freundt, 1983). However, other sera, such as fetal calf serum, may improve the growth potential, and media of quite different formulations have been used satisfactorily for the isolation of both mycoplasmas and ureaplasmas (Shepard, 1983). In particular, SP4 medium, developed originally to cultivate spiroplasmas, has improved the isolation not only of the more fastidious mycoplasmas (TuUy et aL, 1979), but also those more easily isolated, such as M. hominis (Tully et aL, 1983; see also Chapter A2, Vol. I). Pretesting medium components for their ability to support growth and maintaining rigorous quality control, using a fastidious isolate, are important features of successful isolation. Thallous acetate (0.05%), at one time incorporated in medium by most investigators as a bacterial inhibitor, was found to have activity against ureaplasmas (Shepard, 1983) and some mycoplasmas (Tully et aL, 1983) and, therefore, its use has greatly declined. In contrast, the use of one of several synthetic penicillins that have a wide antibacterial spectrum is usual. SPECIMEN PROCESSING
Inoculation of specimens into liquid medium, which is diluted serially, followed by subculture to liquid or agar media provides the most sensitive method for the isolation of many members of the class MoUicutes, including M. hominis and U. urealyticum (Taylor-Robinson, 1983a, 1989). Ureaplasma colonies, in particular, often fail to develop after direct plating of a specimen on agar medium, whereas color changes occur in liquid medium. For the latter to happen, advantage is taken of the metabolic activity of the organisms. Thus, 1.8 ml of medium, supplemented with phenol red (0.002%) and urea (0.1%) and contained
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in a screw-cap vial of 2.5 ml capacity, is inoculated with, for example, 0.2 ml of specimen. Further serial 10-fold dilutions are made as deemed necessary (vide infra), but to at least 10"^. The screw caps are secured and the vials are incubated at 37°C. Ureaplasmas multiply best at pH 6.0 or less and, as they possess a urease that breaks down urea to ammonia, their growth results in the medium, set initially at pH 6.0, changing in color from yellow to pink. In addition, the specimen is diluted in medium containing arginine (0.1%) and, again, in medium containing glucose (0.1%). M. hominis metabolizes arginine by a three-enzyme system via ornithine to ammonia and raises the pH of the medium, set initially at 7.0, so that the color also changes from yellow to pink. Glucose-fermenting mycoplasmas metabolize this substrate to lactic acid by the glycolytic pathway, reducing the pH of the medium, set initially at 7.5-7.8, to less than 7.0, thus changing the color from pink to yellow. The speed of a color change and, therefore, the length of incubation is influenced, at least partially, by the number of organisms in the original specimen. M. genitalium may take months to produce a color change, if it does at all. On the other hand, M. hominis produces a change much more rapidly, usually well within a week, and the ureaplasmas do so usually within 24 to 48 hours or less and rarely thereafter. Serial dilution of specimens, as described earlier, is valuable for various reasons (Taylor-Robinson, 1983a, 1989), most notably in (i) diluting antibodies, antibiotics, and other inhibitors which may be present in the original specimen beyond their inhibitory concentration, and in reducing the number of contaminating bacteria, if present; and (ii) helping to estimate the number of organisms in the original specimen; the highest dilution at which a color change is seen may be regarded as containing one color-changing unit (CCU), assuming that there is even distribution and little aggregation of organisms. As an example, a change up to the 10~5 dilution suggests that the original specimen contained 10^ CCU (or 10^ organisms). To confirm that a mycoplasma has been isolated, aliquots of medium (0.1 or 0.2 ml), taken from cultures which are just changing color or from those which have changed most recently, are introduced into fresh broth medium and/or onto agar medium. In glucose-containing medium at pH 7.5-7.8, minced tissue, cellular deposits, and particularly blood produce a yellow color that may mask any specific change. This demands routine subculture, but otherwise it is usually not profitable to undertake a "blind" passage if color changes have not occurred initially. On agar medium, colonies of the genital mycoplasmas develop best in an atmosphere of 95% N2 plus 5% CO2 (v/v). These mycoplasmas, including M. hominis, produce classical "fried egg" colonies of up to 200 to 300 |xm in diameter. Colonies of M. genitalium are often much smaller and many of these do not have the typical appearance. U. urealyticum organisms produce the smallest colonies (15- to 30-|jLm diameter). Usually these do not have a "fried egg" morphology because they lack the peripheral surface growth. To help to detect
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them, an aliquot of urea-containing broth medium that has changed color should be subcultured to agar medium containing urea, 0.05 M HEPES buffer, and a sensitive indicator of ammonia, either manganous sulfate or calcium chloride (Taylor-Robinson, 1989), on which ureaplasmas form slightly larger dark brown colonies. M. hominis organisms multiply in most routine blood culture media without changing the appearance of the media, and nonhemolytic pinpoint colonies on blood agar following a blind subculture may be the first hint that a patient has had bloodstream invasion. The mycoplasma inhibitory effect of sodium polyethanol sulfonate, included in blood culture media as an anticoagulant, can be overcome by the addition of gelatin (1%, w/v) (Pratt, 1990). Commercial kits designed for the isolation and identification of M. hominis and U. urealyticum are available. Specialist laboratories are likely to have media of superior quality, but successful use of the kits has been reported (Renaudin and Bebear, 1990; Kibsey et al., 1991) and they may be of particular value where the need to detect these microorganisms arises infrequently. Cultured organisms require identification, and certain biological features that are determined routinely during the course of isolation may provide helpful clues. Thus, a specimen from the urogenital tract that produces an alkaline color change without turbidity in medium containing arginine is most likely to contain M. hominis, and one that produces a similar change in medium containing urea is almost certain to contain U. urealyticum. An indication in this way of the mycoplasma that is likely to be present narrows the range of specific antisera that need to be used to make a definitive identification. Incorporation of such antisera in filter paper disks on agar to inhibit colony development (agar growth inhibition) (Clyde, 1964) is widely used, but more than one antiserum to a species may be required to identify various strains of that species (Leach et al., 1987). Furthermore, the close serological relationship between M. genitalium and M. pneumoniae (Lind, 1982) may require several techniques, more than one antiserum (Leach et al., 1987), and perhaps monoclonal antibodies allied to Western blotting (Morrison-Plummer et al., 1987) or immunofluorescence tests (see Chapter B9, this volume) to distinguish between them. Agar growth inhibition has also been used to identify the serotypes of U. urealyticum, although the metabolism-inhibition and complement-dependent mycoplasmacidal tests have been employed more often (Taylor-Robinson, 1983b). Epi-immunofluorescence or immunoperoxidase techniques (Taylor-Robinson, 1983b) used to stain colonies are advantageous in enabling different ureaplasmal serovars or, indeed, mycoplasmal species to be distinguished in mixtures. Antigen or DNA Detection Noncultural procedures have a particular place in the detection of mycoplasmas that are impossible, or almost impossible, to culture (e.g., M. geni-
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talium). Tests to detect antigen have been developed mostly forM. pneumoniae, but irrespective of the mycoplasmal species involved, none has found widespread use because of lack of sensitivity. Inadequate sensitivity also became an obstacle to the development and use of DNA probes. Thus, only 56 and 63%, respectively, of culture-positive urethral specimens from men were found to be positive when tested with probes for M. hominis and U. urealyticum (Roberts et al., 1987). Thus, despite the fact that 30% of urethral specimens from homosexual men proved positive for M. genitalium (Hooton et al., 1988) when tested with a DNA probe which detected as little as 50 to 100 pg of DNA per blot, the general problem of insensitivity has seen the abandonment of straightforward DNA probes and a move to the use of the polymerase chain reaction (PCR) technique by which DNA amplification is induced. DNA primers specific for U. urealyticum (Blanchard et al, 1993) and M. genitalium (Jensen et al., 1991; Palmer et al, 1991), among others, have been developed and used for DNA amplification by the PCR. Details of the procedure relevant to its use for work on mycoplasmas have been presented elsewhere (see Chapter A6, this volume). The method has proved rapid, specific, and sensitive and is of considerable value diagnostically. In the case of M. genitalium, the ability to detect the organisms reliably by means of the PCR, compared to culture, has shown for the first time that they exist primarily in the genitourinary tract and are strongly associated with nongonococcal urethritis (Homer et al., 1993; Jensen ^r a/., 1993). Quantitative Measurement of Antibody Many serological tests have been used to study mycoplasmas (TaylorRobinson, 1989), those of high sensitivity being required for the quantitative measurement of antibody to diagnose infection. However, the extent to which a test is used depends not only on sensitivity but also on specificity, reproducibility, ease of performance, convenience, and familiarity. Lack of sensitivity and specificity make the complement fixation test unsuitable for detecting antibodies to the genital mycoplasmas. The indirect hemagglutination technique is more sensitive and specific and has been used for women with salpingitis to detect antibody responses to M. hominis (Lind et al., 1985) and M. genitalium (Lind and Kristensen, 1987). It is, however, rather complex and difficult to reproduce, and is not used widely. In contrast, the metabolisminhibition test, described in detail previously (Taylor-Robinson, 1983c), is of even greater specificity and has been used to detect antibody responses in various clinical settings, for example, in male subjects after intraurethral inoculation of ureaplasmas experimentally, in those with nongonococcal urethritis and in epididymitis (Mil etai, 1988). The general features and the manner of undertaking enzyme immunoassays (EIAs) have been described previously (Cassell and Brown, 1983). It seems that
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they are becoming more widely used for studying antibody responses to urogenital mycoplasmas. By use of a cell lysate antigen of U. urealyticum and commercially available alkaline phosphatase conjugates, ureaplasmal antibody responses were detected in two-thirds of patients with nongonococcal urethritis (Brown et aL, 1983). Furthermore, a modified EIA has been used to measure changes in the levels of M. hominis antibody classes occurring in women with acute salpingitis (Miettinen et al., 1983). The antigen comprised a soluble cell fraction that masked the serological diversity of different strains and allowed a single representative strain to be used; application of a heavy chain-specific second antibody followed by conjugated antispecies antibodies increased the overall sensitivity. An EIA based on lipid-associated membrane proteins (LAMP) of M. penetrans has been used to measure antibody to this mycoplasma (Wang et aL, 1992) and the same method has also been applied to detecting antibody to M. genitalium. The procedure for the preparation of LAMP and for the EIA has been described in detail (Wang et al, 1992; see also Chapter B4, this volume). The indirect immunofluorescence (IMF) test has gained in popularity. The M. hominis antigen fixed to glass slides was used first in such a test to demonstrate an antibody response in a woman who had a febrile septicemia associated with this mycoplasma (Tully et al, 1965). More recently, an IMF test in which the procedure for measuring antibody to M. genitalium has been described in detail (Furr and Taylor-Robinson, 1984; see also Chapter B7, this volume) was used to detect responses to this mycoplasma in men with nongonococcal urethritis and in women with salpingitis (Taylor-Robinson, 1989). The test is rapid, reproducible, and quite sensitive and specific; there is less cross-reactivity with M. pneumoniae than seen with some other methods.
Discussion The criteria that need to be fulfilled in order to consider that a mycoplasma is an unequivocal cause of a sexually transmitted disease are an extension of Koch's postulates and are applicable to other infectious diseases and other microorganisms. It is clear, however, that it may be difficult or, indeed, impossible to fulfill all the criteria for some diseases. This is particularly so for diseases that involve the upper genital tract. Attempting to determine the effect of a "differential" antibiotic may have to be abandoned since defining one that inhibits the growth of the mycoplasma under consideration but not any other putative etiological agent may prove exceptionally difficult. Furthermore, human experimentation can rarely be expected, if at all. Also, it should be remembered that experiments on human volunteers may give a false impression of what might occur under natural conditions. Although M. hominis organisms, for example.
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produced an exudative pharyngitis when given in large numbers to volunteers (Mufson et al., 1965), no evidence has accrued subsequently to indicate that they are a cause of naturally occurring sore throats. An association between humoral antibody developing after vaccination and resistance to infection and disease and, conversely, lack of antibody correlating with susceptibility to infection and disease are persuasive pieces of evidence that the mycoplasma inducing the antibody is the cause of the disease. However, failure to observe these correlations should not be taken to mean that the mycoplasma is not involved etiologically because vaccination with organisms known to cause a disease does not always provide protection against it. Although many conditions have been associated with urogenital mycoplasmas, only a few are regarded as having a mycoplasmal cause; the mycoplasmas of consequence are M. hominis, M. genitalium, and U. urealyticum. Others, for example, M. fermentans, M. pirum, M. primatum, M. penetrans, M. spermatophilum, and even M. pneumoniae, may cause sexually transmitted disease but an insufficient number of the criteria have been fulfilled, or of attempts made to fulfill them, in order to indicate that this is so. Of course, these criteria and guidelines will need to be invoked for any other mycoplasma that might be discovered in the future. The techniques outlined in this chapter that are required for the detection of mycoplasmas and for the measurement of antibody against them have been presented in detail elsewhere (Taylor-Robinson, 1983a,b,c, 1989, 1994) and have been used to incriminate M. hominis, M. genitalium, and U. urealyticum as causes of disease. Other mycoplasmas should be subjected to the same general procedures. Of these, using an antibody response as the sole criterion for deciding whether infection has occurred or not is unwise, particularly if the decision is based on the existence of antibody in a single serum sample. It should also be noted that antibody responses may develop slowly and that collection of a second "convalescent-phase" serum sample too early could result in failure to detect a rise in the titer of antibody. The slow serological response of male chimpanzees to intraurethral inoculation of M. genitalium emphasizes this point (Tully et al., 1986). Detection of organisms and an antibody response is the ultimate in diagnosis. Detection of an antibody response alone carries less weight than detection of the organisms. In this regard, there is no doubt that the most important recent technical advance has been the development of the PCR technique. This has overshadowed antigen immunoassays and straightforward DNA probes. The PCR technique is invaluable for mycoplasmas that prove difficult or, indeed, impossible to culture. Indeed, it is only by the use of this technique that evidence has accrued to indicate that the culturally difficult M. genitalium is one of the causes of the sexually transmitted disease nongonococcal urethritis (Homer et al., 1993; Jensen et al., 1993). However, isolation of organisms through culture remains a worthy goal since quantitative results are more difficult to achieve with
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the PCR technique and it does not permit an assessment of antibiotic sensitivity or other biological features.
References Blanchard, A., Hentschel, J., Duffy, L., Baldus, K., and Cassell, G. H. (1993). Detection of Ureaplasma urealyticum by polymerase chain reaction in the urogenital tract of adults, in amniotic fluid, and in the respiratory tract of newborns. Clin. Infect. Dis. 17(Suppl. 1), 148153. Brown, M. B., Cassell, G. H., Taylor-Robinson, D., and Shepard, M. C. (1983). Measurement of antibody to Ureaplasma urealyticum by an enzyme-linked immunosorbent assay and detection of antibody responses in patients with nongonococcal urethritis. J. Clin. Microbiol. 17, 288295. Cassell, G. H., and Brown, M. B. (1983). Enzyme-linked immunosorbent assay (ELISA) for detection of anti-mycoplasmal antibody. In "Methods in Mycoplasmology" (S. Razin and J. G. TuUy, eds.). Vol. 1, pp. 457-469. Academic Press, New York. Clyde, W. A. (1964). Mycoplasma species identification based on growth inhibition by specific antisera. J. Immunol. 92, 958-965. Freundt, E. A. (1983). Culture media for classic mycoplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, pp. 127-135. Academic Press, New York. Furr, P. M., and Taylor-Robinson, D. (1984). Microimmunofluorescence technique for detection of antibody Xo Mycoplasma genitalium. J. Clin. Pathol. 37, 1072-1074. Furr, P. M., and Taylor-Robinson, D. (1990). Long-term viability of stored mycoplasmas and ureaplasmas. J. Med. Microbiol. 31, 203-206. Hooton, T. M., Roberts, M. C , Roberts, P. L., Holmes, K. K., Stamm, W. E., and Kenny, G. R. (1988). Prevalence of Mycoplasma genitalium determined by DNA probe in men with urethritis. Lancet i, 266-268. Homer, P. J., Gilroy, C. B., Thomas, B. J., Naidoo, R. O. M., and Taylor-Robinson, D. (1993). Association of Mycoplasma genitalium with acute non-gonococcal urethritis. Lancet 342, 582585. Jalil, N., Doble, A., Gilchrist, C , and Taylor-Robinson, D. (1988). Infection of the epididymis by Ureaplasma urealyticum. Genitourin. Med. 64, 367-368. Jensen, J. S., Uldum, S. A., Sondergard-Andersen, J., Vuust, J., and Lind, K. (1991). Polymerase chain reaction for detection of Mycoplasma genitalium in clinical samples. J. Clin. Microbiol. 29, 46-50. Jensen, J. S., Orsum, R., Dohn, B., Uldum, S., Worm, A.-M., and Lind, K. (1993). Mycoplasma genitalium: A cause of male urethritis? Genitourin. Med. 69, 265-269. Kibsey, P., McKay, T., and Lim-Fong, R. (1991). Rapid detection of Ureaplasma urealyticum and Mycoplasma hominis by Mycofast. Amer. Soc. Microbiol. Abstr. G-13, 135. Leach, R. H., Hales, A., Furr, P. M., Michelmore, D. L., and Taylor-Robinson, D. (1987). Problems in the identification of Mycoplasma pirum isolated from human lymphoblastoid cell cultures. FEMS Microbiol. Lett. 44, 293-297. Lind, K. (1982). Serological cross-reaction between Mycoplasma genitalium and M. pneumoniae. Lancetii, 1158-1159. Lind, K., and Kristensen, G. B. (1987). Significance of antibodies to Mycoplasma genitalium in salpingitis. Eur. J. Clin. Microbiol. 6, 205-207.
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Lind, K., Kristensen, G. B., Bollerup, A. C , Ladehoff, P., Larsen, S., Marushak, A., Rasmussen, P., Rolschau, J., Skoven, I., S0rensen, T., and Lind, I. (1985). Importance of Mycoplasma hominis in acute salpingitis assessed by culture and serological tests. Genitourin. Med. 61, 185-189. Miettinen, A., Paavonen, J., Jansson, E., and Leinikki, P. (1983). Enzyme immunoassay for serum antibody to Mycoplasma hominis in women with acute pelvic inflammatory disease. Sex. Transm. Dis. 10(Suppl.), 289-293. Morrison-Plummer, J., Jones, D. H., Daly, K., Tully, J. G., Taylor-Robinson, D., and Baseman, J. B. (1987). Molecular characterization of Mycoplasma genitalium species-specific and crossreactive determinants: Identification of an immunodominant protein of M.,genitalium. Isr. J. Med. Sci. 23, 453-457. Mufson, M. A., Ludwig, W. M., Purcell, R. H., Gate, T. R., Taylor-Robinson, D., and Chanock, R. M. (1965). Exudative pharyngitis following experimental Mycoplasma hominis type 1 infection. J. Amer. Med. Assoc. 192, 1146-1152. Palmer, H. M., Gilroy, C. B., Furr, P. M., and Taylor-Robinson, D. (1991). Development and evaluation of the polymerase chain reaction to detect Mycoplasma genitalium. FEMS Microbiol. Lett. 61, 199-203. Pratt, B. (1990). Automated blood culture systems: Detection of Mycoplasma hominis in SPScontaining media. In "Recent Advances in Mycoplasmology" (G. Staneck, G. H. Cassell, J. G. Tully, and R. F. Whitcomb, eds.), pp. 778-781. Fischer, Stuttgart. Renaudin, H., and Bebear, C. (1990). Evaluation des systemes Mycoplasma PLUS et SIR Mycoplasma pour la detection quantitative et 1'etude de la sensibilite aux antibiotiques des mycoplasmes genitaux. Pathol. Biol. 38, 431-435. Roberts, M. C., Hooton, M. Stamm, W., Holmes, K. K., and Kenny, G. E. (1987). DNA probes for the detection of mycoplasmas in genital specimens. Isr. J. Med. Sci. 23, 618-620. Shepard, M. C. (1983). Culture media for ureaplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 137-146. Academic Press, New York. Smith, T. F., Weed, L. A., Pettersen, G. R., and Segura, J. W. (1977). Recovery of chlamydia and genital mycoplasma transported in sucrose phosphate buffer and urease color test medium. Health Lab. Sci. 14, 30-34. Taylor-Robinson, D. (1983a). Recovery of mycoplasmas from the genitourinary tract. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 19-26. Academic Press, New York. Taylor-Robinson, D. (1983b). Serological identification of ureaplasmas from humans. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 57-63. Academic Press, New York. Taylor-Robinson, D. (1983c). Metabolism inhibition tests. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, pp. 411-417. Academic Press, New York. Taylor-Robinson, D. (1989). Genital mycoplasma infections. Clin. Lab. Med. 9, 501-523. Taylor-Robinson, D. (1994). Mycoplasmas and ureaplasmas. In "Manual of Clinical Microbiology" (S. Birch and P. Fitzgerald, eds.), 6th ed., pp. 652-662. Am. Soc. Microbiol., Washington, DC. Tully, J. G., Brown, M. S., Sheagren, J. N., Young, V. M., and Wolff, S. M. (1965). Septicemia due to Mycoplasma hominis type 1. A^. Engl. J. Med. 273, 648-650. Tully, J. G., Rose, D. L., Whitcomb, R. F., and Wenzel, R. P. (1979). Enhanced isolation of Mycoplasma pneumoniae from throat washings with a newly modified culture medium. J. Infect. Dis. 139, 478-482. Tully, J. G., Taylor-Robinson, D., Rose, D. L., Furr, P. M., and Hawkins, D. A. (1983). Evaluation of culture media for the recovery of Mycoplasma hominis from the human urogenital tract. Sex. Transm. Dis. 10(Suppl.), 256-260.
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TuUy, J. G., Taylor-Robinson, D., Rose, D. L., Furr, P. M., Graham, C. E., and Barile, M. F. (1986). Urogenital challenge of primate species with Mycoplasma genitalium and characteristics of infection induced in chimpanzees. J. Infect. Dis. 153, 1046-1054. Wang, R. Y.-H., Shih, J. W.-K., Grandinetti, T., Pierce, P. F., Hayes, M. M., Wear, D. J., Alter, H. J., and Lo, S.-C. (1992). High frequency of antibodies to Mycoplasma penetrans in HIVinfected patients. Lancet 340, 1312-1316.
D4 DIAGNOSIS OF NEONATAL INFECTIONS Ken B. Waites and Gail H. Cassell
Introduction Of the 15 species of mycoplasmas isolated from humans, only Mycoplasma hominis and Ureaplasma urealyticum are known to be clinically important in the neonate. Both organisms are a cause of pneumonia, meningitis, bacteremia, and soft tissue abscesses. U. urealyticum has been associated with the development of chronic lung disease of prematurity, death, acute respiratory insufficiency, pneumonia, and persistent pulmonary hypertension of newborns (Cassell and Waites, 1995; CasselUr a/., 1986, 1988, 1993a,b; Waites ^r a/.. 1988, 1993). M. fermentans has been detected in pure culture from placenta and amniotic fluid in the presence of inflammation, thus suggesting it is capable of intrauterine infection. However, no prospective studies to date have been performed to determine its occurrence and significance, if any, in neonates. Although most invasive infections due to M. hominis and U. urealyticum have been described in preterm neonates, those bom at term may also be subject to systemic disease, although apparently less frequently. Routine screening of neonates for M. hominis and U. urealyticum is not indicated. Clinical, radiographic, or laboratory evidence of pneumonia, central nervous system infection, overall instability, or "septic appearance," particularly in a preterm neonate, are reasons to contemplate the possibility of invasive infection with U. urealyticum or M. hominis. Culture techniques are the current diagnostic methods of choice. Consideration of mycoplasmal or ureaplasmal infection is even more important in neonates who have clinical signs and/or symptoms of infection and no other identifiable microbial etiology, or if there is failure to respond to antimicrobial 237 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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treatment with P-lactams and/or aminoglycosides. Cerebrospinal fluid (CSF) mononuclear or polymorphonuclear pleocytosis with a negative gram stain and no growth of bacterial cultures suggests the possibility of mycoplasmal or ureaplasmal infection. However, in some cases in which M. hominis or U. urealyticum is isolated, there is no detectable CSF inflammatory response. Because of the frequency of U. urealyticum colonization of the lower respiratory tract in preterm neonates and its proven association with respiratory disease, culture of endotracheal secretions in all neonates with birth weights of < 1250 g who have respiratory distress at or soon after birth is justified.
Materials Specimens Acceptable specimens for the diagnosis of ureaplasmal pneumonia include endotracheal secretions, pleural fluid, and tissue obtained by lung biopsy. If pneumonia is the primary concern, gastric aspirates, throat swabs, and nasopharyngeal swabs are not recommended since these specimens may not be indicative of the presence of organisms in the lung itself. If other localized or systemic disease is suspected, cultures of CSF, blood, pericardial fluid, urine, or abscess material are appropriate. Transport Media Growth media such as Shepard's lOB broth or SP-4 broth should be inoculated at the time of specimen collection (Waites et al., 1988). Sucrose phosphate transport media (0.2 M sucrose in 0.02 M phosphate buffer, pH 7.2, with 10% heat-inactivated fetal bovine serum), which is widely used for the transport of specimens for chlamydial cultures, is a suitable alternative. It is important to make certain that no antibiotics which could inhibit mycoplasmal growth are present in the transport medium. Failure to inoculate clinical specimens from neonates, most of which will be of very small volume or quantity, into transport media immediately will likely result in drying and possible loss of viability. Growth Media Media for recovery of M. hominis and U. urealyticum from urogenital specimens in adults can also be used for detection of these organisms in neonates. Shepard's lOB broth can be prepared by adding 14 g of commercially available mycoplasma broth base without crystal violet, 2 g of arginine, 0.2 g of DNA, and 1 ml of 1% phenol red to 688 ml ultrapurified water. The pH should be
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adjusted to 5.5 with 2 A^. HCl. The solution is then autoclaved for 15 minutes at 121°C and cooled to room temperature before adding supplements. The following supplements should then be added after each is filter sterlized with a 0.2-|xm filter: 200 ml heat-inactivated bovine serum or non-heat-inactivated horse serum (Hyclone, Logan, UT), 100 ml 25% yeast extract, 5 ml IsoVitaleX [BectonDickinson Microbiology Systems, Cockeysville, MD (BBL)], 4 ml 10% urea, and 2.5 ml 4% L-cysteine. The final pH is adjusted to 5.9-6.1. Penicillin (1000 lU/ml) can be added to suppress growth of contaminating bacteria. The broth may be stored at 4°C for up to 3 months. When arginine is added as described earlier, lOB can be used to cultivate both M. hominis and U. urealyticum. A8 agar is used as the corresponding solid media onto which specimens in lOB broth can be inoculated. A8 agar contains CaCl2 which serves as an indicator for the presence of urease-producing colonies of U. urealyticum. To prepare 1 liter, the following ingredients should be mixed in a flask in the order specified: 825 ml ultrapurified water, 0.15 g CaCl2, 24 g trypticase soy broth (special order 97587, BBL), 2 g yeast extract, 1.7 g putrescine dihydrochloride, 0.2 g DNA, and 10.5 g select agar (BBL). Adjust the pH to 5.5 with 2 A^ HCl and autoclave for 15 minutes at 121°C. Cool in a 56°C water bath. Filter each of the following supplements through a 0.2-|jLm filter separately: 200 ml non-heat-inactivated horse serum (Hyclone), 5 ml IsoVitaleX (BBL), 10 ml 10% urea, 1 ml GHL tripeptide solution (Calbiochem-Novabiochem, La JoUa, CA), and 5 ml 2% L-cysteine. Penicillin (1000 lU/ml) can be added to prevent bacterial contamination. After supplements are mixed with the base, adjust the pH to 6.0, pour plates on a level surface, let sit for 2 hours, invert, and leave overnight. Seal in plastic bags and refrigerate at 4°C for up to 3 months. SP-4 broth and agar were developed for cultivation of spiroplasmas and adapted for isolation of M. pneumoniae. Although both M. hominis and U. urealyticum will grow in SP-4 provided appropriate supplements of arginine and urea are added, its pH of 7.4-7.6 is higher than optimal for U. urealyticum. If both organisms are being sought in clinical specimens, it is most practical to utilize a single broth and agar medium such as lOB broth and A8 agar. Additional Materials Needed
Automatic pipettes and disposable tips to dispense 0.02-0.9 ml Sterile 12 x 75-mm snap-cap tubes Sterile petri dishes Sterile scalpel blades Vortex mixer Stereomicroscope Microscope slides and coverslips Incubators: 37°C, room air atmosphere and 5% CO2 in air atmosphere
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Procedure Specimen Collection Endotracheal secretions can be collected in neonates who are intubated using a small-bore suction catheter attached to a vacuum outlet. The catheter is passed into the trachea through the endotracheal tube and suction is applied. The tip of the suction catheter containing the secretions is then cut with a sterile scalpel blade. One milliliter of lOB broth is drawn into a 3-ml syringe to which a 21gauge needle is attached. The tip of the amputated distal portion of the suction catheter is then placed into the tube from which the lOB broth was drawn. The needle is then used to flush the suction catheter with broth, forcing the endotracheal secretions into the tube which can then be transported to the laboratory. Blood should be collected aseptically without anticoagulants and immediately inoculated into transport medium. A blood:broth ratio of 1:10 with 1 ml blood: 9 ml broth is recommended. Other liquid specimens such as pericardial fluid or cerebrospinal fluid, collected aseptically by pericardiocentesis and lumbar puncture, respectively, should be inoculated into transport medium in a 1:10 ratio, usually a 0.1-ml specimen inoculated into 0.9 ml broth. Urine collected by suprapubic puncture or in a sterile bag attached to the skin around the genitalia can be processed in a similar manner. When sufficient volumes of body fluids such as CSF or urine are available, a portion of the original specimen not inoculated into transport medium should also be sent to the laboratory. Abscess material can be collected using clacium alginate, dacron, or polyester-tipped swabs with plastic or wire shafts. Following collection of the specimen, the swab should be placed into a tube of transport medium, agitated, and excess liquid expressed against the side of the tube. Swabs should then be discarded and not left in the transport medium. Care should be taken to prevent dehydration of tissues obtained by biopsy or autopsy. Tissues can be placed directly into transport medium when collected or, in the case of an autopsy tissue, a carefully excised cube (approximately 1 cm^) can be sent directly to the laboratory in a sterile leak-proof container providing it can be inoculated into culture medium within 1 hour. Specimen Transport All specimens should be transported to the laboratory as soon as possible, even if inoculated into transport media at the time of collection. If immediate transport is not feasible, specimens in transport broth should be refrigerated at 4°C and not left at room temperature. If laboratory processing cannot be accomplished within 6-12 hours following specimen collection or if transportation to a reference laboratory is necessary, specimens in transport medium should be stored at
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—70°C and shipped frozen. Mycoplasmas and ureaplasmas are stable when frozen at — 70°C for long periods of time if maintained in a protein-containing transport medium such as lOB or SP-4 broth. Specimens should never be stored at — 20°C because viability can be lost in a relatively short time. Culture Inoculation and Incubation Frozen specimens should be thawed rapidly in a 37°C water bath. All specimens received in transport media should be thoroughly mixed on a Vortex mixer. Following mixing, specimens should be serially diluted in lOB broth out to 10"^ (0.1 ml original specimen into 0.9 ml broth) using a calibrated pipette so that accurate volumes will be dispensed. If disposable pipette tips are used, the tip should be changed between each transfer to prevent possible carryover which can affect quantitation. A 0.02-ml volume of the original specimen and each dilution should be carefully pipetted onto A8 agar. If agar plates have been refrigerated, it is helpful to place them in an incubator for a short time to allow any condensate to dry since a wet surface can interfere with proper spread of the inoculum and make colony counting difficult. A single agar plate marked into six quadrants can be used for each specimen. After inoculation, the agar plate should be allowed to stand on a level surface until the broth is absorbed prior to placing it into the incubator in an inverted position. If adequate volumes are submitted to the laboratory prior to inoculation into transport medium, liquid specimens such as CSF or urine may be centrifuged at 600 g and cultures as described previously performed on an aliquot from the pellet. Lung tissues from biopsy or tissue collected at autopsy should be removed from transport broth and placed in a sterile petri dish containing a small volume of lOB broth and minced with a sterile scalpel blade prior to being serially diluted. Mincing is preferred to grinding to circumvent the release of potential growth inhibitors. Any portion of an original specimen that remains after all media are inoculated should be frozen at -70°C fof future reference. Broths are incubated at 37°C under atmospheric conditions. Agar plates are incubated in an atmospher of 95% N2 and 5% CO2. The relatively rapid growth rates of M. hominis and U. urealyticum make identification of most positive cultures possible within 2 to 5 days. Growth in lOB broth is suggested by an alkaline shift and a color change from yellow to pink, resulting from either urea hydrolysis by U. urealyticum or arginine hydrolysis by M. hominis. Broths should be examined two to three times daily and tubes showing color change subcultured to agar. Subcultures must be performed soon after a color change occurs, particularly if the organism is U. urealyticum since the culture can lose viability within a few hours. Subculture also increases diagnostic yield since some strains may not grow from the original specimen inoculated initially onto
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solid media. This combination of broth-to-agar inoculation technique has been shown to be the most sensitive for the recovery of mycoplasmas and ureaplasmas from clinical specimens. Cultures should be incubated for at least 7 days before being designated negative. In addition to subculturing to agar, aliquots of broth in which color change has occurred should be frozen in case further studies such as antimicrobial susceptibility testing are needed. If mixed species are present on agar, it will be necessary to pick individual colonies with a swab, inoculate onto a second agar plate, subculture to broth, and freeze for future reference as described earlier. Subculture of colonies on agar to broth can best be accomplished by removing a plug of agar containing colonies, emulsifying in broth, and incubating. Identification of Isolates Agar plates should be examined under a stereomicrosope every 24-72 hours. Mycoplasmal colonies must be distinguished from artifacts such as air bubbles, water or lipid droplets, or other debris which can be confusing. Colonies of U. urealyticum can be identified on A8 agar by urease production in the presence of a CaCl2 indicator following 24-72 hours of incubation. A stereomicroscope is necessary to properly visualize and characterize colonies on A8 agar. U. urealyticum colonies are 15-60 |xm in diameter and appear as brownish granular clumps on the agar surface. Mycoplasmal colonies are approximately 20-300 |xm in diameter, are urease negative, and have the typical "fried egg" appearance on agar. If quantitation is desired, select the dilution which contains wellseparated colonies in numbers that can be readily enumerated. It may be helpful to make a grid on the bottom of the agar plate using a scalpel blade to facilitate accurate counting. The number of organisms in 1 ml of the original specimen is then determined by multiplying the number of colonies times the dilution times 50. Several mycoplasmal species in addition to M. hominis have a similar appearance on agar and no distinguishing biochemical characteristics. The frequency with which other mycoplasmas occur in neonates and any possible associations with disease have not been established. Until species identification can be confirmed, a preliminary report of "large colony Mycoplasma species" is appropriate. Use of growth inhibition by homologous antiserum, immunofluorescence of colonies on agar, immunoblotting with monoclonal antibodies, and the polymerase chain reaction are possible means of speciating mycoplasmas (Blanchard et al., 1993; see also Section A of this volume). Because of the possibility of cross-reactions among species, rigorous proof of the specificity of the method utilized must be documented as well as reactivity of multiple clinical isolates of the same species. None of these methods are readily available as a diagnostic tool for a clinical diagnostic laboratory, making it necessary in most instances to
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submit Mycoplasma isolates to a reference laboratory for complete identification. Definitive evidence that certain serotypes of U. urealyticum are more likely to be associated with disease has not been established. Currently there are no practical methods for serotyping isolates for diagnostic purposes and the procedure is not recommended. Quality Control Attention to all of the procedures outlined for the detection of mycoplasmas cannot guarantee their successful recovery unless extensive quality control of all media components and reagents is practiced. Quality control procedures should include testing newly prepared media with all 14 serotypes of U. urealyticum, the 7 reference strains of M. hominis, and representative low passage clinical isolates. If the number of color-changing units per milliliter between the reference broth and the test broth is more than one to two dilutions lower or if the number of colony-forming units on test agar is less than 90% of the reference agar, the new medium is substandard and should not be used. All new batches should be tested for sterility by incubation at 37°C for at least 24 hours. An uninoculated control tube of complete broth medium should be inoculated with each series of patient specimens to serve as a reference for color change during incubation.
Stams Mycoplasmas in body fluids can be stained with Giemsa, but such preparations may be difficult to interpret due to cellular debris and artifact. A Gram stain may be used to exclude the presence of conventional bacteria. Stains that bind to DNA such as Hoechst 33258 have been used to stain mycoplasmas in normally sterile body fluids, but this technique has not been evaluated in neonates and therefore cannot be recommended as a routine screening method. If it is used, results should always be confirmed by culture.
Discussion As evidence mounts for the pathogenic potential of mycoplasmas and ureaplasmas in a number of clinical conditions that affect neonates, more laboratories will offer diagnostic tests for their detection. In order to achieve valid and reliable results, much attention must be paid to proper specimen handling and quality control due to the sensitivity of mycoplasmas to fluctuating environmental conditions.
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A variety of commercially prepared products designed for the detection of mycoplasmas in diagnostic laboratories are now available. Media sold commercially have yet to be thoroughly evaluated in comparison to the better known nonproprietary formulations that have been in use for several years by research and reference laboratories. Problems with some of the commercially prepared products have included contamination of serum with mycoplasmas of animal origin and media quality control programs limited to testing a single or only a few strains that do not adequately represent low passage clinical isolates and all of the various serotypes and reference strains which may differ in sensitivity to medium components and growth conditions. If commercially prepared media are to be relied on, it is advisable that internal quality control tests be performed prior to use. Serial dilution is an extremely important step in the cultivation process for mycoplasmas as it will help in overcoming possible interference by antibiotics, and other inhibitors that may be present in clinical specimens. Also, when dealing with specimens from nonsterile body sites, dilution helps reduce the numbers of bacteria which could possibly overgrow the mycoplasmas and prevent their detection. Dilution helps overcome the problem of rapid decline in culture viability which is particularly common with ureaplasmas. Alkaline body fluids such as cerebrospinal fluid, blood, and urine may produce a color change in the initial tube of broth. This effect occurs immediately and should not be evident in the serially diluted tubes. Elevated antibodies against M. hominis have been detected in infants with culture-proven meningitis, and the presence of U. urealyticum-specific IgM may be predictive of invasive disease according to some investigators. However, the relative hypogammaglobulinemic state of neonates, especially those bom preterm, the dearth of information concerning the occurrence of antibodies in asymptomatic infants matched for gestational age, the lack of standardized methodology for antibody detection, and the unavailability of commercial reagents make serology a procedure of extremely limited value in the diagnosis of neonatal mycoplasmal infections. Attempts to detect antibodies to M. hominis or U. urealyticum should be limited to specialized research or reference laboratories and cannot be recommended routinely for diagnostic purposes.
References Blanchard, A., Yanez, A., Dybvig, K., Watson, H. L., Griffiths, G., and Cassell, G. H. (1993). Evaluation of intraspecies variation within the 16S rRNA gene of Mycoplasma hominis and detection by polymerase chain reaction. J. Clin. Microbiol. 31, 1358-1361. Cassell, G. H., and Waites, K. B. (1995). Mycoplasmal infections. In "Infectious Diseases of the Fetus and Diseases of the Fetus and Newborn Infant" (J. Remington and J. Klein, eds.), 4th ed., pp. 619-655. Saunders, Philadelphia.
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Cassell, G. H., Clyde, W. A., Kenny, G. E., McCormack, W. M., and Taylor-Robinson, D. (1986). Ureaplasmas of humans with emphasis on maternal and neonatal infections. Pediatr. Infect. Dis. J. 5, S22-S354. Cassell, G. H., Waites, K. B., Crouse, D. T., Rudd, P. T., Canupp, K. C , and Cutter, G. (1988). Association of Ureaplasma urealyticum infection of the lower respiratory tract with chronic lung disease and death in very low birthweight infants. Lancet 2, 240-245. Cassell, G. H., Baseman, J., Lo, S.-C, Quackenbush, R. L., Tully, J. G., Bove, J., Montagnier, L., and Taylor-Robinson, D. (1993a). The changing role of mycoplasmas in respiratory disease and AIDS. Clin. Infect. Dis. 17(Suppl. 1), S1-S315. Cassell, G. H., Waites, K. B., Watson, H. L., Crouse, D. T., and Harasawa, R. (1993b). Ureaplasma urealyticum: Role in prematurity and disease in newborns. Clin. Microbiol. Rev. 6, 69-87. Waites, K. B., Rudd, P. T., Crouse, D. T., Canupp, K. C , Nelson, K. G., Ramsey, C , and Cassell, G. H. (1988). Chronic Ureaplasma urealyticum and Mycoplasma hominis central nervous system infections in preterm infants. Lancet 2, 17-21. Waites, K. B., Crouse, D. T., and Cassell, G. H. (1993). Systemic Ureaplasma urealyticum infections in neonates. Clin. Infect. Dis. 17(Suppl. 1), S131-S135.
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D5 MYCOPLASMAS IN AIDS PATIENTS Shyh-Ching Lo
Introduction Purpose
AIDS (acquired immunodeficiency syndrome) is a unique combination of an infectious process, an autoimmune disorder, and tumorigenesis in afflicted patients. The complex disease often involves multiple organ systems. Functional failure of the immune system leads to various opportunistic infections in these patients. Infection by an agent(s) other than HIV-1 (human immunodeficiency virus type-1) may also be crucial, both in causing a particular illness and in promoting disease progression from nonsymptomatic HIV infection to clinical AIDS. Interest in mycoplasmas has significantly increased because studies from ours and other laboratories showed a much higher frequency of infections with certain unusual species of mycoplasmas in patients with AIDS. The actual role, if any, of these AIDS-associated mycoplasmas is still not clear. But, we have presented three general possibilities for their potential significance in AIDS (Lo, 1992). (A) The mycoplasmas simply represent opportunistic agents found in high frequency in immunocompromised patients with AIDS. (B) Infections with mycoplasmas such as Mycoplasma fermentans and M. penetrans may markedly enhance the pathogenicity of other human viruses including HIV-1, a phenomenon clearly demonstrated in in vitro studies (Lo et ai, 1991b; Lemaitre et al., 1992). Therefore, infections with some species of mycoplasma may promote disease progression to clinical AIDS in HIV-infected patients. (C) The microbe itself is pathogenic in humans. This possibility is supported by our findings in animal experiments using nonhuman primates (Lo et al., 1993d) as well as by the 247 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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association of M. fermentans infection in healthy non-AIDS patients dying of an acute disease (Lo etal, 1989, 1991a, 1993c). It is important to note that mycoplasmal infection is still highly significant clinically in AIDS, even if it is merely another example of an opportunistic infection. It is pathological to have an infectious agent growing in an organ system such as liver, kidney, spleen, lung, or brain. After all, opportunistic infections are the direct cause of death in more than 80% of patients with AIDS. Needless to say, the findings of mycoplasma infection become more significant if the microbe is acting as a disease-promoting "cofactor" or is by itself responsible for certain pathogenic aspects of AIDS. In view of the complex disease process of AIDS, we believe that the mycoplasma may well have a role in all three of the just-mentioned processes. In any case, the unusual microorganisms deserve further study in order to have a better understanding of AIDS. Rationale The pathogenicity or virulence of a microbial agent depends on the precise conditions under which host and infectious agent interact. The infectious organism has to overcome the defensive response of the host in order to gain entry and establish a favorable microenvironment within the host for its continuous survival and replication. Individual conditions of such an interaction between microbes and their human hosts may determine if the infection will have a fulminant course or be completely asymptomatic. Normally, only a portion of individuals who are infected with a microbial agent will actually present a clinical illness. Thus, in a broader term, all clinically significant infections are in a way opportunistic infections. For years before the AIDS epidemic, patients belonging to the high-risk groups of AIDS such as homosexuals, intravenous drug users (IVDU), and hemophiliacs were already known to have different degrees of immune deficiency. They often suffered repetitive infections with various pathogens and were known to be more susceptible to many "less virulent" microbes. Thus, organisms that are otherwise unlikely to grow successfully in healthy human bodies can now colonize in these individuals. Organisms with low pathogenicity may gradually alter the hosts' physiology, cause systemic infection, and produce severe clinical illness. It may also be important to note that for those mycoplasmas which are considered "normal residents" in the human oral cavity and urogenital tract, the status between mycoplasmas and human hosts may have shifted from one of commensalism into one of parasitism and pathogenicity in those individuals with somewhat abnormal immune functions. The human immunodeficiency virus is believed to be the primary cause of AIDS. Infection by HIV may result in permanent damage to the immune system and may eventually lead to even more profound immune deficiency, a condition now termed AIDS.
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Materials and Procedures The techniques used to isolate and identify the AIDS-associated mycoplasmas have been presented in other sections (see Chapter A7 in Vol. I; Chapters B4 and D3, this volume). In general, isolation of fastidious mycoplasmas by culture directly from clinical specimens is still difficult. Polymerase chain reaction (PCR) is a highly sensitive diagnostic technique, but the preparation of clinical samples and each step of the procedure have to be meticulously monitored (see Chapters A4-A8, this volume). The potential complications have kept the technique as an experimental procedure and have prevented most clinical microbiology laboratories from using PCR as a general tool for diagnosis of mycoplasma infections. In addition, one really has to first know where the primary site of infection for the organism occurred before a meaningful assessment of the infection rate or incidence by PCR can be accomplished. Serological assays using ELISA to detect mycoplasma-specific antibodies have been developed (Chapter B4, this volume). This chapter presents a brief summary of mycoplasmas that have been identified much more frequently in patients infected with HIV-1 and/or with AIDS than in non-HIV-infected control subjects. Mycoplasma fermentans
Our laboratory first reported that this organism is most commonly found in the urogenital tracts or urines of patients at high risk of AIDS. Systemic infection with M. fermentans in patients with AIDS is apparently common and can be detected by PCR of urine, blood, diseased tissues, and/or other clinical specimens. Later, others also reported similar findings (Hawkins et al., 1992; Bebear et al., 1993). More importantly, infection by the mycoplasma appeared to be directly associated with functional deficits of the infected organ or tissue. For example, infection by M. fermentans in the kidney is found to be associated with development of AIDS nephropathy in HIV-infected AIDS patients (Bauer et al, 1991). Recently, an HIV-seropositive patient, who was not immunosuppressed, developed focal glomerulosclerosis and M. fermentans DNA was found in the kidney biopsy using PCR. Fifteen months later, M. fermentans was detected in peripheral blood lymphocytes, urine, and throat specimens of this patient. After another 3 months, the CD4+ lymphocyte count had fallen and the patient developed Pneumocystis carinii pneumonia (Ainsworth et al., 1994). This finding strengthens the association between M. fermentans and development of AIDS nephropathy and has illustrated that infection by M. fermentans may precede rapid progression of AIDS. In vitro, the mycoplasma can markedly enhance cytocidal effects of HIV-1 on human CD4+ lymphocytes (Lo et al, 1991b; Lemaitre et al, 1992). Infection by M. fermentans may therefore significantly
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potentiate pathogenicity of HIV-1. The organism is also associated with development of an acute form of adult respiratory distress syndrome which may or may not have systemic involvement in apparently previously healthy individuals (Lo et al., 1989, 1991a, 1993c). Our observations suggest that M. fermentans may cause a previously unrecognized form of human disease which might be systemic, leading to a fatal outcome in previously healthy nonimmunocompromised patients. Mycoplasma genitalium This highly fastidious mycoplasma was previously identified in both urogenital and respiratory tracts and is implicated as a potential pathogen causing pneumonia and nongonococcal urethritis (NGU). Because this is essentially an uncultivable mycoplasma, it has been difficult to assess the scope of infection by this organism without a good seroepidemiological assay. PCR is currently the only diagnostic technique to detect colonization or infection with the organisms. In a PCR study, M. genitalium was found to be associated with development of NGU (Homer et al., 1993). Interest in this mycoplasma increased following a report by Luc Montagnier detecting the organism in blood samples of patients with AIDS by PCR assay (Montagnier et al., 1990). Using a newly developed serological assay, we reported a high frequency of M. genitalium infection in IVDU patients and in male homosexuals with or without HIV infection (Wang et al., 1993). In contrast, very few hemophilic patients with AIDS had serological evidence of M. genitalium infection. Most interestingly, M. genitalium was found to be a rather common infection (>40%) among non-HIV-infected patients attending clinics for treatment of sexually transmitted diseases (STD). Thus, the organism may represent subclinical STD which is not easily recognized or diagnosed. The organism may have a higher chance to produce systemic infection in AIDS patients with compromised immune functions Mycoplasma pirum This organism has an organized terminal structure and can metabolize both glucose and arginine for growth. All previously reported isolates of the species were from human cell cultures. The true origin or natural host for this species has not been well documented. Reports by Luc Montagnier and associates of the isolation of M. pirum from primary cultures of peripheral mononuclear cells prepared from blood of AIDS patients (Montagnier et al., 1990) suggest that M. pirum is a human mycoplasma. This mycoplasma apparently has been isolated on two occasions from the blood of patients with AIDS. However, using ELISA to examine specific antibodies to M. pirum lipid-associated membrane proteins (LAMPs), we found that few patients with AIDS or healthy non-HIV-
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infected individuals had serological evidence of M. pirum infection (Wang et al., 1992). In this context, four isolations of mycoplasmas from 180 urine samples of HIV-1 infected patients with AIDS studied by K. Chirgwin and W. M. McCormack are also believed to be M. pirum. If the true identity of these isolates is confirmed, the uncertainty of the origin of M. pirum can finally be solved. A PCR assay based on the 16S ribosomal RNA gene sequence of M. pirum has been developed (Grau et al, 1993). The highly sensitive assay should be helpful in studying infection by this organism in patients with AIDS. Mycoplasma penetrans This is a previously unknown human mycoplasma (Lo et al., 1992), which our laboratory has isolated more than 12 times from 6 HIV-infected patients. This was done in a study of urine samples obtained from 113 HIV-infected patients. None was isolated from 98 HIV-negative age-matched healthy control individuals in the parallel study. The organism possesses a unique terminal structure by which it attaches and penetrates into eukaryotic cells. Extensive invasion of M. penetrans into infected cells produces cytopathic effects and may lead to cell death. Using electron microscopy, we have directly examined the urine sediments which grew M. penetrans in high titer without any other identifiable infectious agent. M. penetrans could be identified adhering to the surface and invading into the cytoplasm of urothelial cells. M. penetrans with its specialized tip structure apparently uses the same process documented in a variety of cell cultures to adhere onto the cell surface and to invade the cytoplasm of urothelial cells in the urogenital tract of the patient (Lo et al., 1993a). Our preliminary serological study using ELISA and Western blot analyses reveals a high prevalence (40%) of antibodies to M. penetrans in AIDS patients infected with HIV-1. In comparison, a low incidence (0.3%) of antibody is found in sera from non-HIV-infected control subjects. There is also a low prevalence (0.9%) of such antibodies in patients attending sexually transmitted disease clinics. In addition, none of more than 150 HIV-negative patients with different non-AIDS disease states, many associated with immune dysfunction and/or low white cell counts, test positive for the antibodies. The newly discovered mycoplasma is apparently not a commensal organism in humans nor just a simple opportunist commonly occurring in immunocompromised hosts. It is uniquely associated with HIV-1 infection and/or a high risk group(s) of AIDS (Wang et al., 1992). Subsequent serological assays have revealed that the organism primarily infects male homosexuals with or without HIV infection (Wang et al., 1993). Very few HIV-infected patients in other high-risk groups of AIDS are found to be infected by this organism. Since Kaposi's sarcoma occurs mainly in male homosexuals and not in AIDS patients of different risk groups, a significant association can be made between serological evidence of M. penetrans infection
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and development of Kaposi's sarcoma in male homosexuals in the U.S. (/? < 0.001). Thus, the mycoplasma is most likely transmitted through sexual contacts such as anal intercourse among gay men. The newly identified mycoplasma is suspected to have a gastroenteric origin; however, at this time there is no direct evidence to support this speculation. Our original attempts to isolate M. penetrans from rectal swabs of patients with AIDS, including those patients known to be urine culture positive or seropositive for M. penetrans, were not successful.
Discussion Some species of mycoplasmas such as M. fermentans, M. genitalium, M. pirum, and M. penetrans are found in much higher frequency in patients with AIDS or in patients at high risks of having AIDS. Although much has been learned, much more needs to be done before we can reach a conclusion about the actual role of these AIDS-associated mycoplasmas. In studies of various mycoplasmas in HIV-infected individuals, it is interesting to note that isolations of mycoplasmas which are considered "normal residents" such as Ureaplasma urealyticum or M. salivarium from urogenital tracts and the oral cavity are found to be significantly lower in patients with AIDS (Dawson et al, 1993, also Lo et al., unpublished data). One possible explanation is that AIDS patients are often on numerous antibiotics to treat bacterial or parasitic infections. Thus, the commensal mycoplasmas are constantly suppressed. In this case, the opposite findings in frequencies of the AIDS-associated mycoplasmas and of the commensal mycoplasmas in patients with AIDS further suggest that colonization by these AIDS-associated mycoplasmas may be meaningful. Both cross-sectional and prospective cohort studies (Lo et al., 1993b) showed a specific association between infection with M. penetrans and development of Kaposi's sarcoma in HIV-infected male homosexuals, but these highly significant findings of association cannot be considered proof of causality. Even though we do not have complete understanding of the actual role of this mycoplasma, our finding has, however, raised a fundamental question critical in tumor biology. Can persistent infection by mycoplasma(s) induce neoplastic growth of mammalian cells and/or cause cancer in chronically infected hosts like humans? Mycoplasmas may be the only prokaryote which can "symbiotically" grow in the eukaryotic hosts and have a close interaction with mammalian cells for long periods of time. Some mycoplasmas are known to induce a variety of cytokines and have various effects on surfaces of mammalian cells (see Section F in Vol. I). Many potent small proteins of cytokines can effectively mediate a wide range of biological actions on cell proliferation and differentiation in a large spectrum of cells. Intimate interactions on the surface of eukaryotic cells with the microbes may also trigger a cascade of signals transduced from membrane to nuclei.
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altering the functions of many genes. Thus, mycoplasmas may gradually, but significantly, alter a variety of important biological properties of human cells. Since one hallmark of AIDS is the frequent development of many malignancies in the affected young victims, the high frequency finding of some unusual mycoplasmas in patients with AIDS can be significant. The observation of association between infections with M. penetrans and development of Kaposi's sarcoma, a spindle cell tumor, in HIV-infected male homosexuals should prompt further studies of the potential for causing cell transformation by specific species of mycoplasmas.
References Ainsworth, J. G., Katseni, V., Hourshid, S., Waldron, S., Ball, S., Cattell, V., Taylor-Robinson, D. (1994). Mycoplasma fermentans and HIV-associated nephropathy. J. Inf. 29, 323-326. Bauer, F. A., Wear, D. J., Angritt, P., and Lo, S.-C. (1991). Mycoplasma fermentans (incognitus strain) infection in the kidneys of patients with acquired immunodeficiency syndrome and associated nephropathy: A light microscopic, immunohistochemical and ultrastructural study. Hum. Pathol. 22, 63-69. Bebear, C , de Barbeyrac, B., Clerc, M. T., Renaudin, H., Fleury, H. J., Dupon, M., Ragnaud, J. M., and Morlat, P. (1993). Mycoplasmas in HIV-1 seropositive patients. Lancet Ml, 758759. Dawson, M. S., Hayes, M. M., Wang, R.Y.-H., Armstrong, D., Budzko, D. B., Kundsin, R. B., and Lo, S.-C. (1993). Detection and isolation of Mycoplasma fermentans from urine of HIV positive patients with AIDS. Arch. Pathol. Lab. Med. 117, 511-514. Grau, O., Kovacic, R., Griffais, R., and Montagnier, L. (1993). Development of a selective and sensitive polymerase chain reaction assay for the detection of Mycoplasma pirum. FEMS Microbiol. Lett. 106, 327-334. Hawkins, R. E., Rickman, L. S., Vermund, S. H., and Carl, M. (1992). Association of mycoplasma and human immunodeficiency virus infection: Detection of amplified Mycoplasma fermentans DNA in blood. J. Infect. Dis. 165, 581-585. Homer, P. J., Gilroy, C. B., Thomas, B. J., Naidoo, R. O. M., and Taylor-Robinson, D. (1993). Association of Mycoplasma genitalium with acute non-gonococcal urethritis. Lancet 342, 582585. Lemaitre, M., Henin, Y., Destouesse, P., Ferrieux, C , Montagnier, L., and Blanchard, A. (1992). Role of mycoplasma infection in the cytopathic effect induced by human immunodeficiency virus type 1 in infected cell lines. Infec. Immun. 60, 142-14S. Lo, S.-C. (1992). Mycoplasmas and AIDS. In "Mycoplasmas: Molecular Biology and Pathogenesis" (J. Maniloff, R. N. McElhaney, L. R. Finch, and J. B. Baseman, eds.), pp. 525-545. Am. Soc. Microbiol., Washington, DC. Lo, S.-C, Dawson, M. S., Newton, P. B., Sonoda, M. A., Shih, J. W.-K., Engler, W. F., Wang, R. Y.-H., and Wear, D. J. (1989). Association of the virus-like infectious agent originally reported in patients with AIDS with acute fatal disease in previously healthy non-AIDS patients. Am. J. Trop. Med. Hyg. 41, 364-376. Lo, S.-C, Buchholz, C L., Wear, D. J., Hohm, R. C , and Marty, A. M. (1991a). Histopathology and doxycycline treatment in a previously healthy non-AIDS patient systemically infected by Mycoplasma fermentans (incognitus strain). Mod. Pathol. 6, 750-754.
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Lo, S.-C, Tsai, S., Benish, J. R., Shih, J. W.-K., Wear, D. J., and Wong, D. M. (1991b). Enhancement of HIV-1 cytocidal effects in CD4+ lymphocytes by the AIDS-associated mycoplasma. Science 251, 1074-1076. Lo, S.-C, Hayes, M. M., TuUy, J. G., Wang, R. Y.-H., Kotani, H., Pierce, P. F., Rose, D. L., and Shih, J. W.-K. (1992). Mycoplasma penetrans, sp. nov., from the urogenital tract of patients with AIDS. Int. J. Syst. Bacteriol. 42, 357-364. Lx), S.-C, Hayes, M. M., Kotani, H., Pierce, P. F., Wear, D. J., Newton, P. B., Ill, TuUy, J. G., and Shih, J. W.-K. (1993a). Adhesion onto and invasion of mammalian cells by Mycoplasma penetrans—a newly isolated mycoplasma from patients with AIDS. Mod. Pathol. 6, 276-280. Lo, S.-C, Lange, M., Wang, R. Y.-H., Klein, E. B., Inada, Y., Weiss, S. H., and Shih, J. W.-K. (1993b). Development of Kaposi's sarcoma is associated with serologic evidence of Mycoplasma penetrans infection: Retrospective analysis of a prospective cohort study of homosexual men. Abstr. 1st Natl. Conf. Hum. Retroviruses Relat. Infect., Session 67, p. 504. Lo, S.-C, Wear, D. J., Green, S. L., Jones, P. G., and Legier, J. F. (1993c). Aduh respiratory distress syndrome with or without systemic disease associated with infections due to Mycoplasma fermentans. Clin. Infect. Dis. 17 (Suppl. 1) S259-S263. Lo, S.-C, Wear, D. J., Shih, J. W.-K., Wang, R. Y.-H., Newton, P. B., and Rodriguez, J. R. (1993d). Fatal systemic infections of non-human primates by Mycoplasma fermentans (incognitus strain). Clin. Infect. Dis. 17(Suppl. 1), S283-S288. Montagnier, L. (1993). Mycoplasmas as cofactors in AIDS. In "New Concepts in AIDS Pathogenesis" (L. Montagnier and M.-L. Gougeon, eds.), pp. 115-125. Dekker, NY. Wang, R. Y.-H., Shih, J. W.-K., Grandinetti, T., Pierce, P. F., Hayes, M. M., Wear, D. J., Alter, H. J., and Lo, S.-C (1992). High frequency of antibodies to Mycoplasma penetrans in HIVinfected patients. Lancet 340, 1312-1316. Wang, R. Y.-H., Shih, J. W.-K., Weiss, S. H., Grandinetti, T., Pierce, P. F. Lange, M., Alter, H. J., Wear, D. J., Davies, C L., Mayur, R. K., and Lo, S.-C (1993). Mycoplasma penetrans infection in male homosexuals with AIDS: High seroprevalence and association with Kaposi's sarcoma. Clin. Infect. Dis. 17, 724-729.
D6 MYCOPLASMA INFECTIONS OF CATTLE Ed A. ter Laak and H. Louise Ruhnke
Cattle are the hosts of many Mycoplasma species: about 14 Mycoplasma, 3 Acholeplasma, 1 Ureaplasma, and 2 Anaeroplasma species. In addition, Mycoplasma species from other hosts can also infect cattle. Although some mycoplasmas have only limited geographical distribution, a variety of species may occur in one animal or even in one location such as the respiratory or genital tract. Mycoplasma infections can be detected by culture procedures, by indirect methods such as immunological procedures to detect antigens or antibodies, or by detection of mycoplasmal DNA by probes or polymerase chain reaction (PCR). For many bovine mycoplasma species, methods other than culture procedures have not been fully developed or have limitations. Therefore, these methods will be mentioned briefly at the end of this chapter. The isolation methods described in this chapter can also be used with slight modifications for porcine mycoplasmas. The reader is referred to the chapter by Gourlay and Howard (1983) for the characteristics of bovine mycoplasma species. The nutritional demands for isolation of the bovine mycoplasma species differ largely. Various media are needed to isolate bovine mycoplasmas, although most species can be isolated easily on modified Edward's media (medium B of Freundt, 1983). Because only some strains of the strictly anaerobic Anaeroplasma bactoclasticum and A. varium, as well as the facultative anaerobe Mycoplasma alvi, have been isolated and because their significance and prevalence are unknown, these species are not discussed in this chapter. Culture media make use of the different biochemical properties of the various mycoplasma species, in particular their ability to metabolize glucose, arginine, or urea. In this chapter, 255 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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methods for isolation and identification of M. dispar and Ureaplasma diversum will be emphasized because special culture media are required for isolation of these species. M. dispar metabolizes glucose and grows poorly or not at all on solid media on primary isolation. U. diversum metabolizes urea and, although it produces colonies in primary culture, liquid media are also necessary for primary isolation. The media for M. dispar and U. diversum may also allow the growth of other, less fastidious mycoplasma species. Thus, when all possible mycoplasma species are isolated, various types of isolation media and their variants must be used as described under Materials. Because of the considerable international exchange of bovine genetic material and the potential for spread of disease among countries, the protocols for culture of semen are also included.
Materials Anaerobic jars with CO2 and H2 generator envelopes Roller drum (of about one rotation per minute) for tubes Epifluorescence microscope Stomacher 80 Lab-Blender (Seward Laboratory, London) with plastic bags Hanks' minimal essential medium (HMEM) Phosphate-buffered saline solution (PBS), pH 7.2 Rabbit antisera to the various mycoplasma species Antiserum to rabbit immunoglobulins and conjugated to fluorescein isothiocyanate Antiserum to rabbit immunoglobulins and conjugated to peroxidase Nitrocellulose sheet Horse serum: filter sterilized, stored at — 25°C or below in 5- and 10-ml volumes Swine serum: filter sterilized, stored at -25°C or below in 5- and 10-ml volumes Sigma yeast extract: Mix 125 g Sigma yeast type II (YSC-2) into 750 ml distilled water and heat in a water bath 37°C for 25 minutes. Mix frequently. Bring gently to a boil and let it cook for 5-10 minutes. Cool to 56°C and then to 37°C. Centrifuge at 700 g for 20 minutes. Collect the supernatant and dispense in 60-ml volumes. Autoclave (118 to 121°C) for 5 minutes. Store at -25°C or below. Fresh yeast extract (acid extraction): Mix 250 g fresh baker's yeast and 500 ml distilled water at 40°C. Add another 500 ml distilled water at room temperature and mix. Adjust pH to 4.6 with 4 M HCl. Heat to 80°C for 20 minutes. Mix regularly. Cool to 56°C and then to 37°C. Centrifuge at 700 g for 20 minutes. Filter the supernatant through stacked membrane filters, 1.2, 0.8,
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and 0.45 ixm and finally through 0.22 ixm. Dispense in 60-ml volumes. Store at -25°C or below. Fresh yeast extract (aqueous extraction) (modified from Friis and Krogh, 1983): Add 250 g baker's yeast cake to 750 ml distilled deionized water at 80°C, stirring constantly. Heat to 90-100°C for 5 minutes. Cool. Freeze to -20°C. Thaw and centrifuge at 1000 g for 45 minutes. Filter the supernatant through stacked membrane filters, 1.2, 0.8, and 0.45 ixm. Dispense in 10-ml volumes, autoclave for 2-5 minutes, or filter sterilize through 0.22 jxm. Store at -25°C or below. Thallium acetate solution (5.6%): Mix 5.6 g thallium acetate in 100 ml distilled water. Dissolve by heating at 100°C. Dispense in 3-ml volumes. Dispense a 10% solution in 0.2-ml volumes. Store at -25°C or below. Alternatively, the thallium acetate can be autoclaved and stored at 4°C. Phenol red solution (0.5%): Grind 0.5 g phenol red of a water-soluble type in a mortar. Add 15 ml of 0.1 M NaOH little by little during grinding. Add distilled water to a final volume of 100 ml. Store overnight at 4°C. Filter through a 5-|jLm filter. Adjust pH to 7.0. Autoclave for 20 minutes. Dispense in 4.5-ml volumes. Dispense a 1% solution in 0.2-ml volumes. Store at 4°C. Hanks' A: Dissolve 80 g NaCl, 4 g KCl, 1 g MgS04-7H20, and 1 g MgCl2-6H20 in 400 ml distilled water. Add 1.4 g CaCl2 (dry), dissolve completely, and bring volume to 500 ml with distilled water. Heat for 5-10 minutes in a water bath 85°C and dispense in 40-ml volumes. Store at 4°C. Hanks' B: Dissolve 0.745 g Na2HP04-2H20 and 0.6 g KH2PO4 in 500 ml distilled water. Heat for 5-10 minutes in a water bath 85°C and dispense in 40ml volumes. Store at 4°C. Hanks' with dextran: Mix 10 ml Hanks' A, 10 ml Hanks' B, 180 ml distilled water, and 200 mg DEAE-dextran. Dispense in 23-ml volumes. Autoclave for 15 minutes. Store at 4°C. Agar solution: Mix 23 ml Hanks' with dextran (stock solution) and 1.8 g purified agar (Oxoid). Autoclave for 15 minutes. Cool to 56°C. Rabbit hyperimmune serum to M. bovirhinis for SB broth: Mix sera from rabbits, each hyperimmunized with a different strain of M. bovirhinis. In addition to M. bovirhinis type strain PG43, one or more local strains (that are insufficiently inhibited in SB broth containing only antiserum to type strain PG43) may be used. Store at -25°C or below in 10-ml volumes. Diluting solution for antisera in peroxidase test: PBS, pH 7.2, with 0.5 M NaCl, 0.05% Tween 80, and 10% fetal calf serum. Substrate solution for peroxidase test: 12 mg 4-chloro-l-naphthol in 4 ml methanol, 20 ml PBS, pH 7.2, with 12 jxl 30% H2O2, freshly prepared before use. N-medium: Mix 40 ml Hanks' A, 40 ml Hanks' B, 1170 ml distilled water, 5 g brain heart infusion (Difco), and 5.2 g PPLO broth w/o CV (Difco). Autoclave for 15 minutes. Cool to room temperature and add 60 ml Sigma yeast
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extract or fresh acid extracted yeast extract (depending on the culture medium to be prepared), 250 mg bacitracin, 250 mg methicilHn, 4.5 ml 0.5% phenol red solution, and 3 ml 5.6% thallium acetate solution. Dispense in 150- and 10-ml volumes. Store at — 25°C or below. Urea/MgS04 solution: Mix 5 g urea, 2 g MgS04-7H20, and 50 ml distilled water. Bring to the boil in a water bath and let it cook for 5 to 10 minutes. Dispense in 1-ml volumes. Store at -25°C or below. Ureaplasma staining solution: Dissolve 0.25 g urea and 0.2 g MnCl2-4H20 in 25 ml distilled water. Store at 4°C and use within 14 days. Solid and liquid modified Edward's media (see Freundt, 1983) Mycoplasma broth and agar (Truscott and Ruhnke, 1984): Mix 2.1 g PPLO broth w/o CV (Difco) with 70 ml distilled deionized water and 0.2 ml 1% phenol red, and autoclave for 15 minutes. Cool and add sterile supplements: 20 ml inactivated swine serum, 10 ml fresh yeast extract (aqueous extraction), 0.2 ml 50% glucose solution (filter sterilized), 0.5 ml penicillin G potassium 200,000 units/ml, and 0.2 ml 10% thallium acetate solution. Dispense in 1.8ml volumes and store at —25°C or below. For agar, mix 0.8 g purified agar (Oxoid) with the PPLO broth and water, autoclave for 15 minutes, cool to 56°C, and add the same supplements. Dispense in petri dishes, store at 4°C, and use within 7-10 days. NHS25 broth (Medium I of Friis and Krogh, 1983): Mix 150 ml N-medium (including Sigma yeast extract), 25 ml horse serum and 25 ml swine serum (both sera inactivated for 30 minutes at 56°C), and 30 mg cycloserine. Adjust pH to 7.35. Dispense in 1.8-ml volumes. Store at -25°C or below. NHS20 broth (Medium I of Friis and Krogh, 1983): Mix 160 ml N-medium (including Sigma yeast extract), 20 ml horse serum and 20 ml swine serum (both sera inactivated for 30 minutes at 56°C), and 30 mg cycloserine. Adjust pH to 7.35. Prepare NHS agar (see later) or dispense in 1.8-ml volumes. Store the broth tubes at -25°C or below. SB broth (Friis, 1979): Mix 160 ml N-medium (including fresh-yeast extractacid extraction), 20 ml horse serum and 20 ml swine serum (both sera inactivated for 30 minutes at 56°C), and 84 mg cycloserine. Adjust pH to 7.35. Add 10 ml of a mixture of rabbit hyperimmune sera to M. hovirhinis (inactivated for 20 minutes at 56°C). Dispense in 1.8-ml volumes. Store at -25°C or below. NHS agar (Friis and Krogh, 1983): Warm 200 ml NHS20 broth at 56°C for 15 minutes. Add the broth to the sterile molten agar solution that has been cooled to 56°C. Mix well by carefully rotating the bottle so as to avoid air bubbles. Dispense in petri dishes, store at 4°C, and use within 7-10 days. NHU broth, pH 6.0 (Friis et aL, 1980): Mix 160 ml N-medium, 40 ml horse serum (not inactivated), 1 ml urea/MgS04 solution, and 30 mg cycloserine. Adjust pH to 6.0. Dispense in 1.8-ml volumes. Store at -25°C or below.
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Ureaplasma broth: Mix 2.1 g PPLO broth w/o CV (Difco) with 70 ml distilled deionized water and 0.2 ml 1% phenol red solution, adjust pH to 6.0 with 1 M HCl, and autoclave for 15 minutes. Cool and add sterile supplements: 20 ml unheated horse serum, 10 ml fresh yeast extract (aqueous extraction), 1 ml 10% urea solution (filter sterilized), and 0.5 ml penicillin G potassium 200,000 units/ml. Dispense in 1.8-ml volumes. Store at —25°C or below. Ureaplasma agar (modified from Shepard and Lunceford, 1976): Mix 2.1 g trypticase soy broth (BBL, Becton-Dickinson) with 70 ml distilled deionized water and 0.2 ml 1% phenol red solution, adjust pH to 5.5, add 0.8 g purified agar (Oxoid), and autoclave for 10 minutes. Cool to 56°C and add sterile supplements: 20 ml unheated horse serum, 10 ml fresh yeast extract (aqueous extraction), 1 ml 10% urea solution (filter sterilized), 0.5 ml penicillin G potassium 200,000 units/ml, and 0.5 ml 3% MnS04 solution. Dispense in petri dishes, store at 4°C, and use within 7-10 days.
Procedure Preparation of Specimens
Specimens should be processed immediately or stored below -25°C, preferably at -70°C. To prepare inocula, surface sterilize tissue specimens approximately 1-2 cm^ by dipping in boiling water for 6 seconds, then cut off the brown outer surface with sterile scissors and tweezers. Cut about 1 g into small pieces, put into a plastic bag of a Stomacher 80 Lab-Blender together with 2 ml HHEM, and mince for 1 minute. The suspension must be inoculated immediately because lysolecithin released from the host cells will kill the mycoplasmas. This suspension may also be stored at — 70°C for years and can be thawed up to three times without significant loss of numbers of mycoplasmas, as long as it remains unfrozen for only a few minutes. An alternative method is to dip the tissue in ethanol, ignite, and allow to bum off. Cut a clean surface of the tissue with a sterile pair of scissors and rub over)Jthe agar surface. Then mince with scissors in 3 ml broth medium in a sterile petri dish, allow to stand a few minutes, remove the broth to a sterile tube, and make dilutions as described later. Lung or preputial lavage, synovia, or milk can be inoculated directly onto solid and liquid media. Conjunctival, nasal, genital swabs, and mucosal scrapings of the nose may be mixed into 1 ml HHEM before inoculation.
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Isolation of Mycoplasmas
Both solid and liquid media are necessary to recover most of the mycoplasma species. Inoculate solid media by a running drop of the liquid specimens or tissue suspensions. Inoculate liquid media with 0.2 ml of the specimens and make six 10-fold dilutions. Incubate agar media in a CO2 incubator at 37°C for mycoplasmas and at 30° or 37°C for acholeplasmas. Incubate ureaplasma agar anaerobically in CO2 plus H2. Incubate all liquid media aerobically at 37°C for 14 days. Examine agar media for colonies three times per week using a stereomicroscope at 10 to 50 X magnification. Subculture from broth to agar media after 2, 4, and 7 days. ISOLATION OF M. dispar
A lengthy procedure is necessary for isolation of M. dispar because it is a slowgrowing species and is often overgrown by the faster-growing M. bovirhinis, which also metabolizes glucose, or by M. bovis, which does not metabolize glucose, but does acidify the medium. Inoculate 0.2 ml of liquid specimens or tissue suspensions into 1.8 ml NHS25 broth and SB broth and make six 10-fold dilutions. Place the tubes in a roller drum and set to rotate once per minute at 37°C. Observe the broth daily for changes from pink to yellow as the result of metabolism of glucose. Tubes that change color are stored at — 70°C until there are no further changes in the remaining tubes. Subculture the tube (NHS25 broth and/or SB broth) from the highest dilution indicating growth to six dilutions of NHS20 broth and incubate on the drum. Again, observe for acid change and store the broths in -70°C. Inoculate a drop of the highest dilution showing acid pH to NHS agar. Incubate at CO2. Growth of centerless colonies in approximately 1 to 3 days indicates M. dispar. Press three pieces of nitrocellulose sheets gently onto the colonies and remove immediately. Store the pieces at 4°C for 1 week at most until tested by the immunoperoxidase test as described later. ISOLATION OF U. diversum
Inoculate 0.2 ml of liquid specimens or tissue suspensions into 1.8 ml NHU broth, pH 6.0, or ureaplasma broth and make six 10-fold dilutions. Incubate aerobically and without rotation at 37°C. Ureaplasmas produce ammonia, which causes a pH increase in 12 to 18 hours. When the phenol red indicator changes from yellow to light pink, subculture to ureaplasma agar. Incubate anaerobically at 37°C. Examine with a stereomicroscope. Ureaplasmas appear as brown colonies in the presence of MnS04 in 2 to 3 days. If the agar does not contain MnS04, colonies can be identified by flooding the plate with ureaplasmastaining solution and observing for brown colonies after a few minutes. Incubate
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plates for 7 days before discarding as negative. Tubes that are light pink may also be stored at — 70°C for further study. Ureaplasmas are no longer viable when the color of the broth has become purple. ISOLATION OF OTHER MYCOPLASMAS
Modified Edward's broth and agar along with NHS agar may be used for isolation of other mycoplasma species. Broth tubes are incubated aerobically stationary, whereas agar media are incubated in CO2. ISOLATION OF MYCOPLASMAS FROM SEMEN*
The following protocol is recommended to remove antibiotics, which may be only bacteriostatic, used in processing semen (Truscott and Ruhnke, 1984). Mycoplasma broth and agar with ureaplasma broth and ureaplasma agar should be used throughout this procedure. Surface sterilize a semen straw with a cotton gauze soaked in 70% ethanol and let air dry. Aseptically remove semen from straw and place in sterile tube. Plate two (10-|JL1) drops of undiluted semen onto mycoplasma agar and two drops onto ureaplasma agar. Make a 1:10 dilution in mycoplasma broth base (without serum or yeast) in a centrifuge tube (PBS can be inhibitory after 1 hour of exposure). Plate two running drops onto mycoplasma agar and two running drops onto ureaplasma agar. Centrifuge at 35,000 g for 20 minutes at 4°C to 10°C. Discard supernatant and resuspend sediment in 10 ml broth base. Centrifuge at 35,000 g for 20 minutes at 4°C to 10°C. Repeat the washing step twice more. Discard supernatant and resuspend sediment in 1 ml broth base. Plate two running drops onto mycoplasma agar and two running drops onto ureaplasma agar. Make four 10-fold dilutions in ureaplasma broth and one 10-fold dilution in mycoplasma broth. Incubate all broths at 37°C aerobically. Incubate mycoplasma agar in 5-10% CO2 and ureaplasma agar anaerobically. Subculture mycoplasma broth to agar at 2 and 4 (or 5) days. Observe ureaplasma broths daily for color change and subculture to agar as soon as the color changes to light pink. Always confirm a color change by subculture to agar. Examine agar plates every 2 days for mycoplasma and ureaplasma colonies and determine colony count. Incubate plates for 10 days before discarding as negative. Identification of Mycoplasma Species Mycoplasma colonies with a typical fried egg appearance can be identified by the immunofluorescence test (IFT) (Rosendal and Black, 1972; Gardella et al., 1983). Ureaplasmas produce brown colonies on agar containing MnS04 and require no further identification. For serogrouping of U. diversum, which is the * Adapted with permission from Whitford et al. (1994).
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only Ureaplasma species that occurs in cattle, an IFT procedure using calf hyperimmune sera to each of the three serogroups A, B, and C can be applied (Howard and Gourlay, 1981). Most fresh isolates of M. dispar do not produce center-forming colonies and are easily washed off during the IFT procedure. The growth inhibition test is not useful because freshly isolated strains are not or are only slightly inhibited by M. dispar hyperimmune serum. Colonies will attach to the nitrocellulose sheets and can be stained by an immunoperoxidase test using rabbit hyperimmune sera to M. dispar and M. bovirhinis followed by peroxidase-labeled anti-rabbit immunoglobulins (ter Laak and Noordergraaf, 1987). In order to avoid cross-reactions the antisera must be properly diluted with diluting solution. After a final incubation with substrate solution, immunologically stained colonies will appear as purple-blue spots on the nitrocellulose sheets. Other Methods for Detection of Bovine Mycoplasmas Procedures have been reported for the detection of antigens by antigen capture ELISA using monoclonal antibodies to M. bovis (Heller et al., 1993). Small quantities of M. bovis were detected by DNA amplification using polymerase chain reaction. However, direct detection from biological specimens is difficult and requires removal of protein by combined extraction and protease digestion (Hotzel et al,, 1993) (see also Chapter B3, this Volume). Nucleic acid probes for the M. mycoides cluster have been successfully developed for identification of isolates (Taylor et al., 1992a,b). Monoclonal antibodies against M. mycoides subspecies mycoides, small colony biotype, have been characterized and used to improve the diagnosis of contagious bovine pleuropneumonia (Brocchi et al., 1993). Antibody detection by the indirect hemagglutination test has been successful in identifying cattle that have had a previous infection with M. bovis (Cho et al., 1976).
Discussion The media and techniques described earlier are designed to isolate M. dispar and U. diversum. The other bovine mycoplasma species grow easily on solid and in liquid media, such as modified Edward's media. Growth in broth media is indicated by a color change, depending on the glucose- or arginine-degrading characteristics of the species involved. The species M. bovis, M. bovigenitalium, and M. verecundum do not metabolize glucose or arginine and the broth media must be subcultured blindly to agar media. M. dispar grows slowly. A (rapid)
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color change of the NHS25 broth simultaneously in several tubes after 1 to 2 days of incubation often indicates growth of M. bovirhinis. A slower color change after 3 to 5 days, that develops in one tube after the other, indicates growth of M. dispar. When M. bovis is present, this species may also overgrow M. dispar in the liquid media because it grows faster than M. dispar but slower than M. bovirhinis. The growth of M. bovis may be suppressed by adding M. bovis antiserum (final concentration 5%) to the SB broth. The source of CO2 atmosphere is important because commercially available envelope systems that generate a CO2 atmosphere may inhibit mycoplasma growth on agar. Incubators that mix CO2 in the atmosphere and candle jars are usually suitable. The indirect IFT is a convenient and reliable method to identify most mycoplasma species other than M. dispar. However, members of the so-called M. mycoides cluster cross-react in serological identification tests. Species that occur in cattle are M. mycoides subspecies mycoides small-colony biotype and the yet unnamed species Mycoplasma (Leach) bovine group 7. The identification of these species requires additional serologic tests, including the growth inhibition test and the growth precipitation test. It is recommended to send strains suspect of these species to a reference laboratory for proper identification. There are several reasons for diluting the specimens. (1) It is necessary for the isolation procedure of M. dispar. (2) U. diversum and sometimes other mycoplasma species are more often isolated from a liquid culture medium than from a solid culture medium. (3) Often, Ureaplasma has died after overnight incubation in the first dilutions because ureaplasmas grow very fast. By diluting the specimen the ureaplasmas in the more diluted tubes are fewer in number and will survive for a longer period, e.g., 3 days. (4) Mycoplasmas are sometimes inhibited in the first tube (1:10 dilution) in a row of dilutions but do grow in higher dilutions. (5) The number of mycoplasmas in the specimen can be determined and expressed as the number of color-changing units.
References Brocchi, E., Gamba, D., Poumarat, F., Martel, J. L., and De Simone, F. (1993). Improvements in the diagnosis of contagious bovine pleuropneumonia through the use of monoclonal antibodies. Rev. Sci. Tech. Ojf. Int. Epizoot. 12, 559-570. Cho, H. J., Ruhnke, H. L., and Langford, E. V. (1976). The indirect hemagglutination test for the detection of antibodies in cattle naturally infected with mycoplasmas. Can. J. Comp. Med. 40, 20-29. Freundt, E. A. (1983). Culture media for classic mycoplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 127-135. Academic Press, New York. Friis, N. F. (1979). Selective isolation of slowly growing acidifying mycoplasmas from swine and cattle. Acta Vet. Scand. 20, 607-609.
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Friis, N. F., and Krogh, H. V. (1983). Isolation of mycoplasmas from Danish cattle. Nord. Veterinaer med. 35, 74-81. Friis, N. F., Pedersen, K. B., and Bloch, B. (1980). Ureaplasma isolated from the respiratory tract of mink. Acta Vet. Scand. 21, 134-136. Gardella, R. S., Del Giudice, R. A., and Tully, J. G. (1983). Immunofluorescence. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, pp. 431-439. Academic Press, New York. Gourlay, R. N., and Howard, C. J. (1983). Recovery and identification of bovine mycoplasmas. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 81-89. Academic Press, New York. Heller, M., Berthold, E., Pfutzner, H., Leirer, R., and Sachse, K. (1993). Antigen capture ELISA using a monoclonal antibody for the detection of Mycoplasma bovis in milk. Vet. Microbiol. 37, 127-133. Hotzel, H., Demuth, B., Sachse, K., Pflitsch, A., and Pfutzner, H. (1993). Detection oi Mycoplasma bovis using in vitro deoxyribonucleic acid amplification. Rev. Sci. Tech. Off. Int. Epizoot. 12, 581-591. Howard, C. J., and Gourley, R. N. (1981). Identification of ureaplasmas from cattle using antisera prepared in gnotobiotic calves. J. Gen. Microbiol. 126, 365-369. Rosendal, S., and Black, F. T. (1972). Direct and indirect immunofluorescence of unfixed and fixed mycoplasma colonies. Acta Pathol. Microbiol. Scand., Sect. B 80, 615-622. Shepard, M. C , and Lunceford, C D . (1976). Differential agar medium (A7) for identification of Ureaplasma urealyticum (human T mycoplasmas) in primary cultures of clinical material. J. Clin. Microbiol. 3, 613-625. Taylor, T. K., Bashiruddin, J. B., and Gould, A. R. (1992a). Application of a diagnostic DNA probe for the differentiation of the two types of Mycoplasma mycoides subspecies mycoides. Res. Vet. Sci. 53, 154-159. Taylor, T. K., Bashiruddin, J. B., and Gould, A. R. (1992b). Relationships between members of the Mycoplasma mycoides cluster as shown by DNA probes and sequence analysis. Int. J. Syst. Bacteriol. 42, 593-601. ter Laak, E. A., and Noordergraaf, J. H. (1987). An improved method for the identification of Mycoplasma dispar. Vet. Microbiol. 14, 25-31. Truscott, R. B., and Ruhnke, H. L. (1984). Control of mycoplasma and ureaplasma in semen. Proc. Annu. Meet.—Am. Assoc. Vet. Lab. Diagn., SuppL, pp. 50-64. Whitford, H. W., Rosenbusch, R. F., and Lauerman, L. H., eds. (1994)."Mycoplasmosis in Animals: Laboratory Diagnosis." Iowa State Univ. Press, Ames.
D7 MYCOPLASMA INFECTIONS OF GOATS AND SHEEP A. J. DaMassa General Introduction Goats and sheep are important commodities to a large segment of the world's population as a source of meat, milk, and fiber, sometimes as companion animals, and, in certain instances, they have been and continue to be used as sacrificial offerings in ritualistic religious services. Clinical mycoplasmosis often lacks pathognomonic characteristics, and symptoms can be shared by or can mimic other clinical infections. In a previous document by Cottew (1983), which remains relevant, the basic methodology for the diagnosis of mycoplasma infections in these hosts was set forward. This addendum is intended to briefly outline only those aspects that require updating. Some of these changes include: (a) new or altered species names, (b) occurrence of mycoplasma in the ear canal of lactating goat breeds, (c) oral infection of the newborn by ingestion of the mycoplasma in the mammary secretions, and (d) infection of the lactating doe by the introduction of the agent into the teat canal during milking.
Nomenclatural Changes and New Species Several new nomenclatural changes have been proposed for mycoplasmas within this group (Table I). These include the creation of two subspecies of Mycoplasma capricolum: M. capricolum subsp. capricolumsind M. capricolum subsp. capripneumoniae (Leach et aL, 1993). Additionally, three new species 265 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
266
A. J. DaMassa TABLE I BIOCHEMICAL CHARACTERISTICS OF MYCOPLASMA ISOLATED FROM GOATS AND SHEEP"
Mycoplasma M. adleri M. agalactiae M. arginini M. auris M. bovirhinis M. bovis M. capricolum subsp. capricolum M. capricolum subsp. capripneumoniae M. conjunctivae M. cottewii M. gallinarum M. mycoides subsp. capri M. mycoides subsp. mycoides (LC) M. ovipneumoniae M. putrefaciens M. yeatsii M. sp. 2D A. axanthum A. granularum A. laidlawii A. oculi Ureaplasma spp.
Type strain G145 PG2 G230 UIA PG43 Donnetta Calif, kid F38 HRC581 VIS PG16 PG3 Y Y98 KS-1 GIH 2D S743 BTS39 PG8 19L
G
A
T
P
I
F
D
+ + + + + + + + +
+ + + -
+ +
+ + +
+
+ -
+ + +
S
+ -
+ + + +
S
+ V
NT^ NT
V
-f
+ + + + + + + +
V
-
W
w
-1-
+ + +
+^ + + -
+ -
+ + -
NT
NT
NT
NT
s
+ + -f
W
-
-f
+ + + -f-
+ + + + + + + + + NT
''G, glucose; A, arginine; T, tetrazolium (aerobic); P, phosphatase; I, digestion of inspissated serum; F, film and spots formation; D, digitonin; V, variable reaction; S, slow, positive reaction; W, weak positive reaction; LC, large colony or caprine biotype; M., Mycoplasma; A., Acholeplasma. *Most, if not all, large colony isolates are phosphatase positive (rarely negative) if the determination is made by broth procedures; most are negative if tested by agar plate tests (even if plates are incubated for 3 weeks). All of the 43 United States isolates tested in this laboratory by a broth procedure were phosphatase positive. '"Not tested.
from the external ear canal of goats have been proposed, M. auris, M. cottewii, and M. yeatsii (DaMassa et al., 1994), and the name M. adleri has been proposed for a Strain (G145) isolated from a caprine foot (Del Giudice et al., 1995).
Characteristics of Caprine-Ovine Mycoplasmas Tables I, II, III, and IV provide current knowledge of the biochemical, pathological, usual isolation sites, and probable contagiousness of this group of mycoplasmas.
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TABLE II PATHOGENIC MYCOPLASMA OF GOATS AND SHEEP AND USUAL DISEASE PROCESSES WITH WHICH THEY ARE ASSOCIATED^
Mycoplasma/type strain
Process Affects sheep (unusual, except with Ma, Mcon) Arthritis/polyarthritis Celluhtis Conjunctivitis Lung lesions Adhesions to costal wall Fibrinous pleuritis Distended interiobular septa Mastitis Pyrexia Septicemia
MA PG2
Mc CK
Mcc F38
Mmc PG3
Mmm Y
Mp KSl
My GIH
Y98
+
+
-
7
+
-
-
+
+ +
+ + +/-
-
+ + 7
+ + +
+/-
-
+
+/-
+ + + -
+/+/-
+/+/-
+ -
+ -
-
+ +
+/-
+/-
+/+ + +
7 4-
+ +
+/+ +
+
MO
^Results are those normally encountered in lactating goat breeds, except forM. capricolum subsp. capripneumoniae; disease attributable to this species is normally encountered in African goats, where arthritis and mastitis are not normally seen. Ma, M. agalactiae; Mc, M. capricolum subsp. capricolum; Mcc, M. capricolum subsp. capripneumoniae; Mmc, M. mycoides subsp. capri; Mmm, M. mycoides subsp. mycoides (LC); Mp, M. putrefaciens; My, M. yeatsii; Mo, M. ovipneumoniae. +/-, positive to negative (variable response); CK, California kid.
Occurrence of Mycoplasmas in Ear Canal of Goats An important development in caprine-ovine mycoplasmology occurred in 1981-1982 with the disclosure of the occurrence of mycoplasmas, often several pathogenic species, in the external ear canal of lactating goat breeds. These findings have now been verified on several occasions, although similar results from African goat breeds have thus far not been demonstrated. Mycoplasmas in the external ear constitute an important finding because this site represents the most probable one for the demonstration of mycoplasmas in the clinically normal, live, lactating goat. Often, the mycoplasmas are found in close association with ear mites, which some investigators believe are agents that transfer mycoplasmas to the new hosts that mites colonize. It is highly probable that the presence of mycoplasmas in the ear canal may not occur (except in cases of septicemia) in herds without a history of mycoplasma infection or when mites are absent. As many as six species and about 10^ colony-forming units (CPUs) of mycoplasma have been demonstrated in a single ear swab culture. Mycoplasmas are particularly prevalent in goats throughout the world; the association between
268
A. J. DaMassa TABLE III
USUAL ANATOMICAL ISOLATION SITES OF COMMON MYCOPLASMAS OF GOATS AND SHEEP
Mycoplasma
Isolation site
M. agalactiae" M. arginini M. auris M. bovis M. cottewii M. capricolum subsp. capricolum^ M. capricolum subsp. capripneumoniae M. conjunctivae M. mycoides subsp. capri^ M. mycoides subsp. mycoides'^ M. ovipneumoniae M. putrefaciens^ M. yeatsii"^ M. sp. 2D Acholeplasma oculi Ureaplasma spp.
Mouth,^ udder, joints, external ear, eyes, vagina Mouth, respiratory tract,^ eyes External ear Lung External ear Mouth, udder, joints, respiratory tract, external ear Respiratory tract Eyes, respiratory tract Mouth, udder, joints, respiratory tract, external ear Mouth, udder, joints, respiratory tract, external ear, eyes Mouth, respiratory tract, eyes Udder, joints, external ear Mouth, lung, udder, external ear Genital tract Eyes Genital tract
«Host may become septicemic; in such cases, mycoplasma isolations are possible from many sites, including blood. ^In the mouth, the tonsillar crypts provide a rich cultural site. <^Includes lung lesions, mediastinal lymph nodes, and pleural fluid. ^Results based on a single report in which the organism was most likely septicemic.
TABLE IV CAPRINE MYCOPLASMOSIS: TRANSMISSION THROUGH CONTACT EXPOSURE BY PATHOGENIC MYCOPLASMAS^
Contact transmission among Mycoplasma M. M. M. M. M. M.
agalactiae capricolum subsp. capricolum capricolum subsp. capripneumoniae mycoides subsp. capri mycoides subsp. mycoides putrefaciens
Adults
Kids
Low (?) Low High Low (?) Low Low
Low (?) High High High (?) High Low
"With all mycoplasmas listed except M. mycoides subsp. capri, where insufficient data exist, recorded evidence indicates that close contact is a requirement for transmission to occur.
07 Mycoplasma Infections of Goats and Sheep
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mycoplasmas and ear mites (particularly Raillietia caprae) may be an important factor. The presence of mycoplasmas in the ear canal of sheep has not been demonstrated.
Culture of External Ear Canal for Mycoplasmas and Mites Materials 13 X 100-mm (or other convenient size) screw-capped, clear glass tubes containing 3-5 ml mycoplasma medium "B" (see "Media Formulations") or other suitable medium. Sterile cotton-tipped applicators on wooden or plastic sticks (overall length about 150 mm) (American Scientific Products, McGaw Park, IL).
Procedure 1. Two operators are normally required, one to hold the animal and the other to culture the ear. The pinna of the ear is held upward with a gentle pull in a plane parallel to the face. A swab is inserted downward into the orifice of the ear canal and pushed inward about 3-5 cm until a slight resistance is encountered. A few rotations of the swab completes the process. The swab is withdrawn and inserted into the screw-capped tube just past the cotton bud; the bud is broken or cut off and placed into the media. 2. The glass tubes containing the medium and swabs are examined under a stereomicroscope with a magnification of 15-20X for ear mites. The medium must be examined for mites within 1-3 hours of culturing, before incubation at 37°C. Incubation imparts turbidity to the medium, thereby obscuring visibility. 3. Two species of ear mites (ArachnidaiAcari) are normally encountered in the external ears of domestic goats: Psoroptes cuniculi (AcariiPsoroptidae) and Raillietia caprae (Acari:Mesostigmata). Mites have three pairs of legs as larva and four pairs as nymphs and adults. Psoroptes mites have a characteristic psoroptiform outline, with the first two pairs of legs (adults) situated anteriorly and projecting mostly forward and the third and fourth pairs situated and pointing mostly backwards behind the idiosoma (body). On further magnification, each leg terminates in a long, segmented stalk. R. caprae has the general body outline of ticks, with the four pair of legs (adults) spaced evenly apart and distributed along the sides of the body. 4. The liquid medium is plated onto the corresponding solid medium after 2 and 5 days incubation at 37°C.
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A. J. DaMassa
5. Colonial growth is examined for morphologically different mycoplasmas, which are isolated and identified. If no morphologic differences are noted, an arbitrary number of colonies (6-10) should be examined. Generally, the ear-swabbing technique has severe limitations as a measure of ear mite prevalence. P. cuniculi can be demonstrated with more regularity than R. caprae by the swab technique, probably because they are more superficial, i.e., nearer the external entrance to the ear canal. Usually, the external ear canal contains waxy deposits, and in many cases ear mites are congregated between a thick plug of wax and the tympanum, beyond the reach of swabs. Raillietia spp. are more apt to be obscured by this waxy plug. Ear mites can be demonstrated with more regularity by otoscopic examination than by culture.
Oral Infection of Newborn through Colostrum or Milk Lactating goats infected with several different mycoplasmas often shed large numbers of the agent, up to 10^ or more CFUs/ml of colostrum or milk. With some strains, young kids rapidly acquire the infection through ingestion of the mycoplasmas in the mammary secretions. This method of transmission can often be avoided by (1) removing the kid from its dam immediately after birth, before nursing, and (2) feeding either a mixture of heat-treated colostrum (minimum 50%) and milk (56°C for 30 minutes), or (less preferably) milk replacer.
Media Formulations The media formulations and methodology for biochemical tests previously given (Cottew, 1983) remain satisfactory and should continue to be used. Additional, useful formulations follow: 1. Yeast extract, prepared according to a procedure described previously (Freundt, 1983) and used at a final concentration of 10-15% is satisfactory for caprine-ovine mycoplasmas. 2. Solid and liquid medium "B" (Freundt, 1983) is satisfactory for most caprine-ovine mycoplasmas. 3. FF medium (Freundt, 1983) is useful for the growth of M. ovipneumoniae. 4. For M. capricolum subsp. capripneumoniae, two media formulations are in use: (a) special purpose liquid and solid medium prepared from viande foie digest (Cottew, 1983), and (b) medium "WJ" (Jones and Wood, 1988), which is prepared as follows. Autoclavable portion: Bacto-PPLO broth (without CV; Difco Laboratories,
D7 Mycoplasma Infections of Goats and Sheep
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Detroit, MI), 1.5 g; Bacto-tryptone (Difco), 1.5 g; Bacto-peptone (Difco), 0.3 g; Bacto-yeast extract (Difco), 0.1 g; distilled water, 50 ml (broth) or 55 ml (agar); agarose (Miles Scientific, Naperville, IL), 0.9 g (agar). Autoclave for 15 minutes at 120°C. Add the following membrane-filtered (220 nm) components: newborn calf serum (inactivated, 56°C for 30 minutes), 45 ml (broth) or 40 ml (agar); tissue culture medium 199 (10x, without sodium bicarbonate, with glutamine), 5 ml; fresh yeast extract, 5 ml; 0.2% calf thymus DNA, highly polymerized (Sigma Chemical Co, St. Louis, MO), 1.0 ml; 50% glucose, 0.5 ml; 10% nicotinamide adenine dinucleotide (Sigma), reduced form (NADH), 0.1 ml; 10% thallous acetate, 0.25 ml; ampicillin (100 mg/ml), 0.25 ml; 0.4% phenol red indicator, 1.5 ml. Final pH is 7.6-7.8.
Cultural Sites for Mycoplasmas 1. From the live animal: The preferred cultural sites are eyes, genital tract, joint aspirates in case of arthritis, milk, mouth, nares, oropharynx, and the external ear canal. Blood is an important cultural site, particularly if pyrexia is present (Note: some mycoplasmas do not elicit pyrexia). If the animal can be sedated, the tonsillar crypts should be cultured. 2. From the animal at necropsy: These include the blood, brain, eye, genital tract, joint fluid, liver, lung, lymph nodes (mediastinal, supramammary), spleen, udder, and tonsillar crypts.
Milking Procedures The udder is a prime target organ for mycoplasma infection. Small numbers of mycoplasmas belonging to several species can initiate mastitis leading to overt disease if introduced into the teat canal. This is most often caused by unsanitary procedures (improper disinfection), either in the milking parlor or by hand milking. Management practices that are helpful in curtailing this type of transmission follow: 1. A sample of milk from each "bulk tank" should be cultured for mycoplasma. A mycoplasma-positive bulk tank sample requires that the infected animals be identified. This is most easily accomplished by dividing the animals into successive smaller groups or "strings." The milk of each smaller group can be pooled and cultured until the mycoplasma-positive animals are identified. 2. Mycoplasma-positive animals should be isolated into a mycoplasma-
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A. J. DaMassa
positive "string" and milked last. Culling of mycoplasma-positive animals should be considered. 3. Udders should be cleansed, disinfected, and dried before milking, whether the goats are milked by milking machines or are hand milked. The teat cups of the milking machines should be disinfected often, particularly if mastitis occurs in the herd. Milking machines that prevent "backflush" are desirable.
Note on Mycoplasma Presence Many clinically normal goats, particularly lactating breeds, may harbor several mycoplasmas in diverse anatomical sites. Examples are clinically normal herds that have a herd incidence of 80% or more of mycoplasmas (sometimes several pathogenic species) in the ear canal. Pathogenic mycoplasmas can also be encountered in the mouth and respiratory tract of the clinically normal goat. Their isolation from a joint or internal organ is, however, always cause for concern. These principles apply to mycoplasmas exclusive of M. capricolum subsp. capripneumoniae, where insufficient data exist.
Comparison of Pathological Data In describing the gross or microanatomic changes attributable to a specific mycoplasma, there is a tendency to assume that the response is the same in all goats, which is not always the case. As an example, M. mycoides subsp. mycoides (LC) has been known to be a prime initiator of arthritis in lactating goat breeds, although it does not seem to elicit those manifestations in African goats. This discrepancy has been apparently verified (Adesotoye and Ojo, 1990) by the demonstration that strains of M. mycoides subsp. mycoides known to cause arthritis in North American goats do not elicit that manifestation in Nigerian goats.
References Adetosoye, A. I., and Ojo, M. O. (1990). Failure to induce experimental arthritis in Nigerian goats by Mycoplasma mycoides subsp. mycoides, LC type. lOM Lett. 1, 343-344. Cottew, G. S. (1983). Recovery and identification of caprine and ovine mycoplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 2, pp. 91-104. Academic Press, New York. DaMassa, A. J., Tully, J. G., Rose, D. L., Pitcher, D., Leach, R. H., and Cottew, G. S. (1994).
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Mycoplasma auris, sp. nov., M. cottewii sp. nov., andM. yeatsii sp. nov., new sterol-requiring mollicutes from the external ear canal of goats. Int. J. Syst. Bacteriol. 44, 479-484. Del Giudice, R. A., Rose, D. L., and Tully, J. G. (1995). Mycoplasma adleri, sp. nov., an isolate from a goat. Int. J. Syst. Bacteriol. 45, 29-31. Freundt, E. A. ( 1983). Culture media for classic mycoplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 2, pp. 127-135. Academic Press, New York. Jones, G. E., and Wood, A. R. (1988). Microbiological and serological studies on caprine pneumonias in Oman. Res. Vet. Sci. 44, 125-131. Leach, R. H., Emo, H., and MacOwan, K. J. (1993). Proposal for designation of F38-type caprine mycoplasmas as Mycoplasma capricolum subsp. capripneumoniae subsp. nov. and consequent obligatory relegation of strains currently classified as M. capricolum (Tully, Barile, Edward., Theodore and Emo 1974) to an additional new subspecies, M. capricolum subsp. capricolum subsp. nov. Int. J. Syst. Bacteriol. 43, 603-605.
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D8 MYCOPLASMA INFECTIONS OF SWINE Richard F. Ross and Gerald W. Stemke
Introduction The major pathogenic mycoplasma in swine is Mycoplasma hyopneumoniae, a widespread cause of pneumonia. Two other pathogens in swine include M. hyorhinis, a cause of polyserositis and arthritis in 3- to 10-week-old pigs, and M. hyosynoviae, a cause of arthritis in pigs 10 to 24 weeks of age (Ross, 1992). Additional information on experimental models for the induction of M. hyopneumoniae disease and the nature of the disease is presented in Chapter E6 of this volume. Information on methods for isolation of porcine mycoplasmas was presented by Ross and Whittlestone (1983); relatively little has changed with respect to media or approaches utilized for recovery of these organisms during the intervening years. Methods for isolation of M. hyopneumoniae, given in detail in Ross and Whittlestone (1983), included media formulations, selective procedures for isolation of the organism in the presence of M. hyorhinis, collection and specimen preparation, incubation, observation and subcultivation, and traditional identification procedures. Certain aspects of the diagnosis of these diseases were reviewed also by Ross (1993). Specific diagnosis of M. hyopneumoniae disease may be accomplished through use of immunofluorescence, immunoperoxidase, DNA probe, and polymerase chain reaction (PCR) methods for detection of the organism in the swine lung. Examples of these various methods for diagnosis of M. hyopneumoniae infection are presented in this chapter. The enzyme-linked immunosorbent assay (ELISA), procedure is most often used at present for serodiagnosis of M. hyo275 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. 11
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Richard F. Ross and Gerald W. Stemke
pneumoniae disease; therefore, approaches to use of the ELISA are also described.
Immunofluorescence Procedures We have successfully used immunofluorescence for direct detection of M. hyopneumoniae in porcine lung. Either a direct (Amanfu et al., 1984) or an indirect (Piffer and Ross, 1985) procedure can be utilized. Lung tissue should be collected from obviously pneumonic areas; lung specimens for sectioning should transect bronchi and bronchioles in the affected area. These specimens should be embedded in the appropriate medium for frozen section procedure, such as OCT compound (Amanfu et al., 1984). Sections may be cut at 4 jxm with a cryostatmicrotome, fixed for 10 minutes in absolute methanol at 4°C, air dried, and stored at 20°C. For the direct immunofluorescence procedure, a fluorescein isothiocyanate (FITC)- conjugated M. hyopneumoniae antibody (porcine) is prepared according to conventional procedures. Sections mounted on slides are coated with the appropriately diluted conjugate in 0.01 M phosphate-buffered saline (PBS) (pH 7.4) and incubated in a moist chamber for 30 minutes. The uncombined conjugate is removed by rinsing the slides twice with precooled PBS for 5 minutes for each rinse with stirring. Sections are counterstained with a chelated eriochrome black T counterstain for 30 seconds and washed twice in precooled distilled water for 2 minutes for each rinse. The air-dried sections are mounted in phosphate-buffered glycerol (pH 7.4) and observed with a fluorescence microscope. Positive specimens reveal a fluorescent coating of M. hyopneumoniae antigen on bronchial and bronchiolar epithelial surfaces. The strongest fluorescence is observed during acute and midstage disease with a decrease during later stages of disease. It is important that immunofluorescence be utilized with lungs that have been collected from animals that have been killed or died very recently; the process of autolysis proceeds rapidly so that the epithelium sloughs from the airways and the organisms cannot be detected in close apposition to the ciliated epithelium. Lungs collected from swine with chronic M. hyopneumoniae lesions at slaughter or during late stage disease are generally immunofluorescence negative.
Immunoperoxidase Procedures Several reports have appeared in which M. hyopneumoniae has been detected in pneumonic lungs using an enzyme-linked immunoperoxidase technique (Bruggmann et al., 1971 \ Doster and Lin, 1988). The procedure, described by
D8 Mycoplasma Infections of Swine
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Doster and Lin (1988), is an adaptation of that reported by Hill (1978). Lung specimens fixed in 10% neutral-buffered formalin are dehydrated through a graded series of ethanol and xylene, embedded in paraffin, sectioned at 2 jxm, and mounted on slides. Porcine origin antibodies against M. hyopneumoniae are used as the primary antibody to coat tissue sections, and peroxidase-conjugated rabbit antibody to swine IgG (Cooper Biomedical, Inc., Malvern, PA) diluted 1:640 is used as a linking antibody. Observation with a light microscope reveals the mycoplasmal organisms as pleomorphic dark-brown granules (0.3 to 0.5 |xm in diameter) coating the bronchial and bronchiolar epithelium of affected tissues. Doster and Lin (1988) commented that the distribution of the organisms was similar to that seen with the immunofluorescence procedure. The advantage of the immunoperoxidase procedure is that it can be utilized with formalinpreserved tissues.
Gene Probes and PCR Techniques DNA probes to detect and characterize mycoplasmas isolated from pig lungs have been reported by several workers (Stemke, 1989; Ahrens and Friis, 1991; Abiven et aL, 1992; Johansson et al., 1992; Futo et al, 1992). Although these are useful for unambiguous identification of cultured mycoplasmas, direct detection in clinical materials from chronically infected pigs has not been demonstrated because of limitations of sensitivity. However, Ahrens and Friis (1991) and Johansson et al. (1992) have detected M. hyopneumoniae in clinical specimens from some acutely infected animals using their probes. Several investigators have developed PCR for the detection and identification of M. hyopneumoniae of porcine mycoplasmas isolated from pig lung tissues (Harasawa et al, 1991; Artiushin et al, 1993; Stemke et al, 1994). Although direct detection by PCR from clinical samples using these methods has not been reported, unambiguous identification of the organisms in small volume cultures can be achieved. We have utilized nested PCR using primers and conditions reported for universal mollicute 16S rRNA PCR (Deng et al, 1992) for the first stage and then used specific porcine respiratory mycoplasmal primers binding well within the 16S rRNA gene to yield species-specific small PCR products (Stemke et al, 1994) for the nested stage. A few microliters of the amplified material of the first stage is directly used for the second stage without purification, but with approximately five times the primer concentration of the first stage. This modification allows the detection of the DNA from less than 10 organisms. Although the first stage amplification is not restricted to mycoplasmas, the nested primers are specific under the conditions employed. Using this method we have detected M. hyopneumoniae in pig lung samples from
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Richard F. Ross and Gerald W. Stemke
animals that were not clinically ill, i.e., apparently carrier animals. If this method proves generally applicable, it may make detection much more practical than presently is the case.
Serodiagnostic Techniques Serodiagnostic tests utilized for detection of M. hyopneumoniae have included the indirect hemagglutination test, the complement fixation test, and various formats of ELISA (Ross and Whittlestone, 1983). Current evidence indicates that ELISA is the most useful because of sensitivity, specificity, and duration of antibody responses (see Chapter B3, this volume). Nicolet et al. (1980) developed an ELISA based on use of a Tween 20 extract of M. hyopneumoniae. Evaluation by other workers has indicated that this ELISA offers substantial improvement over tests described previously (Bereiter et al,, 1990; Bolske et al., 1990; Kazama et al, 1989). Others have used the Tween 20 ELISA to detect antibodies in both the serum and the colostrum of sows from infected herds. In 1992, Feld et al. reported the development of a blocking ELISA based on use of a biotinylated monoclonal antibody against a 74-kDa polypeptide in M. hyopneumoniae. The ELISA, developed by Nicolet et al. (1980), is performed using a Tween 20 extract of M. hyopneumoniae cells as antigen as described in Chapter B3 in this volume. Modifications used in our laboratory are described by Bereiter et al. (1990). Specifically, polystyrene microtiter plates are Immunolon 2 plates (Dynatech Laboratories, Inc., Alexandria, VA), wells in plates are coated with 100 |xl of 10 |JLg/ml of Tween 20 Ag in carbonate-bicarbonate buffer (pH 9.6), and plates are incubated overnight at room temperature and stored at -70°C. Washing is done as described in chapter B3, except that we use PBS containing 0.05% Tween instead of 0.01% Tween 20. One hundred microliters of test sera, strongly positive control sera (absorbance 0.6-0.7), weakly positive control sera (absorbance 0.3-0.4), and negative control sera (absorbance 0.1-0.2) is diluted 1:50 in Tris-buffered saline solution containing 0.1% bovine serum albumin, 1 mM EDTA, and 0.05% Tween 20 (TBS dilution buffer) and added to duplicate wells on the plate. Plates are then incubated at 37°C for 30 minutes. The peroxidase conjugate is added as in Chapter B3 in this volume and incubation is carried out at 37°C for 30 minutes. The wells are washed again, and 100 |xl of hydrogen peroxide and ABTS (2,2'-azinodi[3-ethylbenzthiazoline sulfonate(6)]) (Kirkegaard and Perry Laboratories Inc., Gaithersburg, MD) is added and incubated at 37°C for 10 minutes. The reaction is stopped by adding 100 JJLI per well of 1% sodium dodecyl sulfate, and the net absorbance values (absorbance of test sample minus absorbance of wells without serum) are measured with an auto-
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mated microplate reader at 405 nm (Model AL310; Bio Tek Instruments, Inc., Winooski, VT). Sera having absorbance values greater than 0.26 are considered positive. Absorbance values between 0.20 and 0.26 are suspect. Absorbance values less than 0.20 are considered negative. The blocking ELISA described by Feld et al. (1992) is carried out using the supemate of sonicated M. hyopneumoniae cells as the antigen according to the following steps. 1. The wells of microtiter plates (Nunc, Roskilde, Denmark) are coated with 100 |xl per well of rabbit immunoglobulin to M. hyopneumoniae diluted 1:1000 in 0.1 M carbonate buffer, pH 9.6. 2. Plates are incubated overnight at 4°C, tapped dry, and blocking is carried out for 1 hour with a solution of 4% skim milk in PBS (designated PBS-M). 3. Washing is done four times with PBS containing 2% Tween 20 (designated PBS-T) and this is followed by a 1-hour incubation at room temperature with the M. hyopneumoniae antigen diluted 1:250 in PBS-T. 4. Plates are washed again, and test sera, diluted 1:10 in PBS-M, are added in duplicate and the plates are incubated for 2 hours. 5. A biotinylated MAb against a M. hyopneumoniae epitope of approximately 74 kDa, diluted 1:2000 in PBS-M, is added to the wells without emptying the antigen mixture and these are incubated for 15 minutes at room temperature. conjugated avidin (Dako, Glostrup, Denmark) diluted 1:8000 in PBS-T. 7. Plates are washed again and an enzyme substrate (8 mg 1,2-50% inhibition are considered positive.
Summary The diagnosis of M. hyopneumoniae disease in the field requires a combination of approaches, including clinical evaluation of the swine herd for evidence of signs of the disease, postmortem examination of selected individuals for the
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detection of typical gross lesions of the disease, and histopathologic evaluation for evidence of typical microscopic changes induced by M. hyopneumoniae. As described in this chapter, more precise methods for specific detection of M. hyopneumoniae in swine tissue may include use of immunofluorescence, immunoperoxidase, DNA probe, or PCR techniques. Several modifications of the ELISA procedure have been shown to provide the specificity and sensitivity needed for the detection of antibodies to M. hyopneumoniae in swine sera. Methods used for the diagnosis of M. hyopneumoniae disease vary substantially from case to case and depend on the objectives of the clinical veterinarian and the needs of the individual swine producer.
References Abiven, P., Blanchard, B., Saillard, C , Kobisch, M., and Bove, J. M. (1992). A specific DNA probe for detecting Mycoplasma hyopneumoniae in experimentally infected piglets. Mol. Cell. Probes 6, 423-429. Ahrens, P., and Friis, N. F. (1991). Identification of Mycoplasma hyopneumoniae DNA with a DNA probe. Lett. Appl. Microbiol. 12, 249-253. Amanfu, W., Weng, C. N., Ross, R. P., and Barnes, H. J. (1984). Diagnosis of mycoplasmal pneumonia of swine: Sequential study by direct immunofluorescence. Am. J. Vet. Res. 45, 1349-1352. Artiushin, S., Stipkovits, L., and Minion, F. C. (1993). Development of polymerase chain reaction primers to detect Mycoplasma hyopneumoniae. Mol. Cell. Probes 7, 381-385. Bereiter, M., Young, T. P., Joo, H. S., and Ross, R. P. (1990). Evaluation of the ELISA and comparison to the complement fixation test and radial immunodiffusion enzyme assay for detection of antibodies against Mycoplasma hyopneumoniae in swine serum. Vet. Microbiol. 25, 177-192. Bolske, G., Johansson, K.-E., Strandberg, M.-L., and Bergstrom, K. (1990). Comparison of the cross-reactions to different Mycoplasma hyopneumoniae antigen preparations in ELISA. Zentralbl. BakterioL, Suppl. 20, 832-834. Bruggmann, S., Engberg, B., and Ehrensperger, P. (1977). Demonstration of M. suipneumoniae in pig lungs by the enzyme-linked immunoperoxidase technique. Vet. Rec. 101, 137. Deng, S., Hiruki, C , Robertson, J. A., and Stemke, G. W. (1992). Detection by PCR and differentiation by restriction fragment length polymorphism of Acholeplasma, Spiroplasma, Mycoplasma, and Ureaplasma, based upon 16S rRNA genes. PCR Methods Appl. 1, 202-204. Doster, A. R., and Lin, B. C. (1988). Identification of Mycoplasma hyopneumoniae in formalinfixed porcine lung, using an indirect immunoperoxidase method. Am. J. Vet. Res. 49, 17191721. Peld, N. C , Qvist, P., Ahrens, P., Friis, N. P., and Meyling, A. (1992). A monoclonal blocking ELISA detecting serum antibodies to Mycoplasma hyopneumoniae. Vet. Microbiol. 30, 35-46. Futo, S., Seto, Y., Mitsuse, S., and Mori, Y. (1992). Detection of Mycoplasma hyopneumoniae by using rRNA-oligonucleotide hybridization. J. Clin. Microbiol. 30, 1509-1513. Harasawa, R., Koshimizu, K., Takeda, O., Uemori, T., Asada, K., and Kato, I. (1991). Detection of Mycoplasma hyopneumoniae DNA by the polymerase chain reaction. Mol. Cell. Probes 5, 103-109.
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Hill, A. C. (1978). Demonstration of mycoplasmas in tissues by the immunoperoxidase technique. J. Infect. Dis. 137, 152-154. Johansson, K.-E., Mattsson, J. G., Jacobsson, K., Fernandez, C , Bergstrom, K., Bolske, G., and Wallgren, P. (1992). Specificity of oligonucleotide probes complementary to evolutionarily variable regions of 16S rRNA from Mycoplasma hyopneumoniae and Mycoplasma hyorhinis. Res. Vet. Sci. 52, 195-204. Kazama, S., Yagihashi, T., and Seto, K. (1989). Preparation of Mycoplasma hyopneumoniae antigen for the enzyme-linked immunosorbent assay. Can. J. Vet. Res. 53, 176-181. Nicolet, J., Paroz, P., and Bruggmann, S. (1980). Tween 20 soluble proteins of Mycoplasma hyopneumoniae as antigen for an enzyme linked immunosorbent assay. Res. Vet. Sci. 29, 305309. Piffer, I. A., and Ross, R. F. (1985). Immunofluorescence technique for detection of Mycoplasma hyopneumoniae in swine lungs. Pesqui. Agropecu. Bras. 20, 877-882. Ross, R. F. (1992). Mycoplasmal diseases. In "Diseases of Swine" (A. D. Leman, B. E. Straw, W. L. Mengeling, S. D'Allaire, and D. J. Taylor, eds), 7th ed., pp. 537-551. Iowa State Univ. Press, Ames. Ross, R. F. (1993). Mycoplasma—animal pathogens. FEMS Symp. 62, 69-109. Ross, R. F., and Whittlestone, P. (1983). Recovery of, identification of, and serological response to porcine mycoplasmas. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 115-127. Academic Press, New York. Stemke, G. W. (1989). A gene probe to dQiect Mycoplasma hyopneumoniae, the etiological agent of enzootic porcine pneumonia. Mol. Cell. Probes 3, 225-232. Stemke, G. W., Phan, R., Young, T. F., and Ross, R. F. (1994). Differentiation of Mycoplasma hyopneumoniae, M. flocculare, and M. hyorhinis on the basis of amplification of a 16S rRNA gene sequence. Am. J. Vet. Res. 55, 81-84.
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D9 MYCOPLASMA INFECTIONS OF POULTRY Stanley H. Kleven and Sharon Levisohn
Numerous mycoplasma species have been isolated from one or more species of poultry or other birds (see Table I). Diagnosis of mycoplasmosis in poultry is usually directed toward detection of infection by the major pathogenic species: Mycoplasma gallisepticum, M. synoviae, M. meleagridis, and M. iowae. Host specificity among the avian mycoplasmas is fairly strict: M. gallisepticum and M. synoviae are pathogenic for both chickens and turkeys and while isolated from other types of birds, they are considered to be pathogenic primarily in gallinaceous species. M. gallisepticum has been isolated from house finches (Carpodacus mexicanus) in the eastern United States. Putative isolations of M. gallisepticum from goose, duck, and partridge were found to constitute a new species, M. imitans. M. meleagridis is confined to turkeys, whereas M. iowae infects both turkeys and chickens but has been shown to be naturally pathogenic only in turkey embryos. These mycoplasmas are sometimes isolated from other avian species, but it is not clear to what degree this reflects passive transmission or genuine colonization. Nonetheless, such findings can have major significance for epidemiological studies and control programs. Pigeons have a distinct mycoplasma flora (Table I), for which the pathogenicity properties have not been fully clarified. M. anseris, M. cloacale, and an unnamed species designated 1220 have been isolated and associated with field syndromes in geese, including embryo mortality, poor growth, egg transmission, infection of the cloaca and phallus, and infertility (Stipkovits et al., 1987). The designation of four new mycoplasma species (Table I), each isolated from a different species of raptor, suggests that isolation attempts in other avian species may be rewarding. This chapter addresses the general principles of isolation and identification of 283 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 19% by Academic Press, Inc. All rights of reproduction in any form reserved.
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TABLE I BIOCHEMICAL CHARACTERISTICS OF AVIAN MOLLICUTES"
Mycoplasma species
Type strain
Usual host
Mycoplasma anatis M. anseris M. buteonis M. cloacale M. columbinasale M. columbinum M. columborale M. corogypsi M. falconis M. gallinarum M. gallinaceum M. gallisepticum M. gallopavonis M. glycophilum M. gypis M. imitans M. iners M. iowae M. lipofaciens M. meleagridis M. pullorum M. synoviae Ureaplasma gallorale
1340 1219 Bb/T2g 383 694 MMPl MMP4 BVl H/Tl PG16 DD PG31 WRl 486 Bl/Tl 4229 PG30 695 R171 17529 CKK WVU1853 D6-1
Duck Goose Buzzard Various Pigeon Pigeon Pigeon Black vulture Saker falcon Chicken Chicken Chicken, turkey Turkey Chicken Griffon vulture Duck, goose, partridge Chicken Turkey Chicken Turkey Chicken Chicken, turkey Chicken
Glucose fermentation
+ + + + + + + + + + + + + —
Arginine hydrolysis
Phosphatase activity
+ -
+ -
-f
+ + + + + + + + + —
-h
)
+ + 7
"Adapted from Jordan (1983), with permission.
avian mycoplasmas, including sample collection and processing. Several, or even many, different Mycoplasma species may be isolated from the same bird, and diagnostic procedures are usually designed to focus on the pathogen(s) and to identify them. However, some of the so-called saprophytic species may have at least limited pathogenic capability, especially in debilitated animals or in mixed infection with other pathogenic agents (viruses or bacteria) or with other mycoplasmas. The use of gene-based methods such as gene probes and polymerase chain reaction (PCR) for identification of all the pathogenic species is relatively highly developed and provides a realistic alternative or supplementary diagnostic method for some types of clinical samples, which will be discussed. Since all of the pathogenic species are egg transmitted, control programs for maintaining breeder flocks free of pathogenic mycoplasmas are important. An essential part of control programs is methodology for obtaining a rapid diagnosis
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of infection in infected flocks, even in the absence of clinical signs. Serological testing is the basis for mycoplasma control programs, and methods for the detection of antibodies to the avian species will be outlined.
Recovery and Identification of Avian Mycoplasmas Growth Medium General principles for the cultivation of mycoplasmas are also applicable for avian mycoplasmas. No single medium formulation has been universally accepted as optimum for growth of all the species. Table II shows examples of two commonly used formulations for media which support the growth of all avian mycoplasmas (Frey et al., 1968; Jordan, 1983). Medium for the isolation of ureaplasma and specific suggestions for preparation of medium components have been described (Jordan, 1983). A yeast source may be beneficial and is usually supplied by commercial yeast autolysate or a fresh yeast extract. M. synoviae uniquely requires nicotinamide adenine dinucleotide (NAD), and cysteine hydrochloride is added as a reducing agent for the NAD. In general, horse serum should be used in media for M. meleagridis, swine serum for M. synoviae, and either horse or swine serum for M. gallisepticum and M. iowae. Some laboratories use a mixture of the two types of sera. The substitution of ampicillin (200-1000 mg/liter) for penicillin G may be made in agar and broth media; the inhibitory spectrum of ampicillin is broader, and there are generally somewhat fewer problems with bacterial contamination. Thallium acetate can be added up to a final concentration of 500 mg/liter. Glucose is fermented by M. gallisepticum, M. synoviae, M. iowae, and several other species and is a common supplement. When used together with phenol red as a pH indicator, color change may be a useful indicator of growth. Sample Collection During the acute stages of infection with M. gallisepticum, M. synoviae, or M. meleagridis (generally in the first 1-2 months postinfection), the population of organisms in the upper respiratory tract and the incidence of infection in the flock are high. In such cases, swabbing of the trachea or choanal cleft of the live bird (10-20 samples) is usually sufficient, whereas as many as 30-100 cultures may be required at later stages. For M. iowae or M. meleagridis it may be useful to culture the cloaca, phallus, vagina, or semen of turkeys. Excretion of mycoplasmas and levels of organisms found in these samples vary greatly.
TABLE II FORMULATIONS FOR TWO COMMONLY USED MEDIA FOR ISOLATION AND PROPAGATION OF AVIAN MYCOPLASMAS
Medium
Constituent
Amount
Prey's
Mycoplasma broth base (BBL, Cockeysville, MD) Glucose Swine serum Cysteine hydrochloride'^ Nicotinamide adenine dinucleotide (NAD)" Phenol red (1%) Thallium acetate (10%)* Penicillin G potassium*^ Distilled water q.s. Adjust pH to 7.8 with 20% NaOH and filter sterilize^-^
22.5 g 3g 120 ml 0.1 g 0.1 g 2.5 ml 2.5 to 5 ml 106 units 1000 ml
PPLO brotb/"
PPLO broth without crystal violet (Difco) Glucose Swine serum Fresh yeast extract^ Cysteine hydrochloride" NAD" Phenol red (1%) Thallium acetate (10%)* Penicillin G potassium'^ Distilled water q.s. Adjust pH to 7.8 with 20% NaOH and filter sterilize^-^
14.7 g 10 g 150 ml 100 ml 0.1 g 0.1 g 2.5 ml 2.5 to 5 ml 106 units 1000 ml
"Reduced NAD is required for A/, synoviae only. A 1% solution each of NAD and cysteine is mixed in equal parts, and 20 ml is added per liter of medium. *For potentially contaminated specimens, use 5 ml of 10% thallium acetate per liter, and add an extra 20 ml of 1 % thallium acetate per liter of medium to bring total concentration to 1:1500. Add thallium acetate to the distilled H2O before the other ingredients to prevent precipitation of protein. "^For potentially contaminated material, an extra 2 x 10^ units of penicillin may be added per liter of medium; 200 mg to 1 g of ampicillin per liter of medium will substitute. ^Alternatively, all ingredients except cysteine, hydrochloride/NAD, serum, penicillin, and yeast extract may be autoclaved at 1 2 r c for 15 minutes, and the remaining ingredients are added aseptically after sterilization by filtration. ^ Fresh yeast extract (also available commercially) is made by placing 250 g dry baker's or brewers' yeast in 1 liter of distilled H2O and allowing to soak for 1 hour. Heat to boiling, allow to cool, and centrifuge at 3000 g for 20 minutes. Decant the supernatant fluid and adjust the pH of the fluid to 8.0 with O.IM NaOH. Clarify by filtration through coarse filter paper and sterilize by filtration. Dispense in aliquots and store at -20°C. ^For agar medium, use 1% of a purified agar such as ion agar no. 2, Noble agar, or Difco purified agar. All components except cysteine, hydrochloride/NAD, serum, and penicillin are sterilized by autoclaving at 121°C for 15 minutes. Cool to 50°C and aseptically add the above components that have been sterilized by filtration and warmed to 50°C. Mix and pour plates to a depth of approximately 5mm. Note: From Kleven and Yoder (1989). A Laboratory Manual for the Isolation and Identification of Avian Pathogens by American Association of Avian Pathogens. Copyright 1989 by American Association of Avian Pathogens. Reprinted with permission of Kendall/Hunt Publishing Company.
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When airsacculitis, synovitis, or other lesions indicative of mycoplasmosis are present, the lesion in necropsied birds should be cultured, usually by swabbing the surface of the affected organ. In some cases, culture of other tissues such as oviduct, intestinal tract, or brain is recommended, even in the absence of pathological changes (Jordan and Amin, 1980). Organisms tend to disappear from the lesions after a few weeks, although usually persisting at least at a low level in the respiratory tract. Secondary infections with other bacteria are also more prevalent at later stages of infection. Therefore, isolation from sacrificed birds at the early stage of disease will probably be more successful than isolation attempts from dead birds or individuals with chronic infection. For culture from embryonated eggs or dead embryos samples of yolk, including a portion of the yolk membrane or swabbing from the yolk sac membrane at late stages of incubation (18 days for chicken eggs or a corresponding period in other birds) is recommended. Culture of the esophagus or yolk sac of the embryo in pipped eggs is often successful for isolation of the pathogenic mycoplasmas. Mycoplasmas may also be isolated from yolk membrane of nonfertile eggs. When specimens must be stored or shipped to a laboratory, it may be useful to inoculate broth medium on the farm and ship by overnight carrier to the laboratory. Inoculated broth cultures are viable for several days at room temperature, although they should be incubated as soon as possible. Alternatively, tracheas, fluids, or tissues can be collected and frozen on dry ice. Isolation of mycoplasmas is generally quite successful when specimens are preserved on dry ice and arrive at the laboratory still frozen. Shipment on wet ice may be necessary in some cases, but specimens should be cultured within 24 hours, if possible. Procedure
Samples (swab or tissue) are usually inoculated into the broth medium of choice; the swab should then be discarded to reduce the possibility of the bacterial contamination. Some investigators prefer to inoculate one or more serial dilutions to avoid inhibitory effects of inoculum components such as lytic enzymes or residual antibiotic. M. gallisepticum and especially M. synoviae cultures are very sensitive to low pH, and broth cultures should be subcultured and/or plated as soon as a color change is detected in medium containing the phenol red indicator. In the absence of color change (as for nonfermenters or in medium without glucose), cultures are plated at about 5 days of incubation and again at 10-14 days, using a cotton swab or a Pasteur pipette. Agar plates can be divided into sections so that as many as eight broth cultures can be inoculated on a single agar plate. Agar plates are examined microscopically for mycoplasma colonies. With several of the nonpathogenic species, such SLS Acholeplasma laidlawii, M. gallinarum, or M. gallinaceum, colonies may be observed as early as 24 hours postinoculation. However, with the pathogenic species, M. gallisepticum,
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M. synoviae, and M. meleagridis, colonies are most frequently observed at 4-5 days of incubation at 37°C, but in some instances they may be present as early as 3 days. M. iowae isolates are very variable, with some types forming large rapidly growing colonies and others requiring at least a week for colony detection. Strain variation is also encountered in the other species, and some isolates are very slow growing. Thus evidence of growth may not be observed until 2 to 3 weeks of incubation. Mixed cultures are common, especially in adult chickens. The most commonly encountered nonpathogenic species are M. gallinarum and M. gallinaceum. Direct plating to agar may allow slow-growing colonies of the potential pathogens M. gallisepticum and M. synoviae to be distinguished from rapidly growing large colony saprophytes which would overgrow in broth culture. The addition of hyperimmune serum against contaminating mycoplasmas (20 ml per liter of medium) may retard their growth enough to improve the possibility of detecting the pathogenic species. Elimination of the NAD supplement from the medium may facilitate the isolation of M. gallisepticum from mixed infection with M. synoviae. For M. meleagridis and M. iowae, primary isolation directly on agar plates may be more successful than inoculating broth medium. Identification of Isolates Identification of avian mycoplasmas is carried out by standard methods. Biochemical properties are not of great value since pathogenic strains M. gallisepticum and M. synoviae have the same properties, as do several of the saprophytes (Table I). Isolates must be cloned in order to carry out these tests. However, certain properties may be expressed even in uncloned isolates. Film and spot production is characteristic of M. synoviae, although there is strain variation and it may not always be distinguishable on all growth media. Phosphatase production is a property of several species, the most frequently found of which is M. meleagridis. When the test is carried out on agar medium (Bradbury, 1983) it may be used to detect the presence of M. meleagridis even when this is present as a minor component in mixed cultures. Direct or indirect immunofluorescence of mycoplasma colonies (Gardella et al., 1983) is a rapid, reliable procedure which can be used with mixed cultures. This procedure is routinely used to identify field isolates in many laboratories. Using visible light to locate nonfluorescing colonies in mixed cultures and then switching to a UV light source can be useful for identifying single fluorescing colonies in mixed cultures. The use of conjugates against a second mycoplasma species employing a different fluorescent dye may also be useful for rapidly identifying mixed cultures. Immunoperoxidase procedures, either alone or in combination with immunofluorescence, may also be useful for rapid identification of mycoplasma cultures (Bencina and Bradbury, 1992). Growth inhibition with specific hyperimmune antiserum (Clyde, 1983) re-
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mains one of the most reliable means of identifying isolates and can be recommended for laboratories that do not have access to a fluorescent microscope. However, it is necessary to begin with a pure culture, and several days may be required to complete the procedure. For any of the just described methods it is necessary to have a set of hyperimmune sera produced in rabbits or avian species (see Chapter E5, this volume).
Serological Tests for Avian Mycoplasmas Diagnosis of mycoplasma infections in poultry breeder flocks is often carried out in the absence of overt clinical signs. Screening of the flock for infection with M. gallisepticum and M. synoviae is generally accomplished with the serum plate agglutination (SPA) test. Positive reactors in the SPA test are then confirmed by further serological testing and/or demonstration of the presence of the organism by culture or gene-based methods. Serological testing is also used as a laboratory diagnostic aid for clinical samples and in experimental trials. An example of a control program is published in the provisions of the United States Department of Agriculture, National Poultry Improvement Plan (NPIP) (1993). Generally, 10% of the flock (or a minimum of 300 birds) is tested before the onset of egg production, and approximately 30 birds per flock are tested every 60-90 days thereafter. A diagnosis is made on a flock basis, and the presence of one or more individual infected birds constitutes an infected flock. Serum Plate Agglutination Test The SPA test is quick, inexpensive, and highly sensitive. Specific stained antigens for M. gallisepticum, M. synoviae, and M. meleagridis are available commercially (Intervet, Boxmeer, Holland) and are used according to manufacturer's instructions. Infected birds may test positive as early as 7-10 days after infection, but occasionally agglutination tests are insensitive, especially for the detection of M. synoviae antibodies in turkeys. The greatest advantage of the SPA test is low specificity (false-positive reactions). Some of these may be related to medium components, primarily serum, adhering to the surface of the mycoplasma organisms used to prepare the antigen. False-positive reactions related to medium components are often seen after vaccination against other infectious diseases, although in other cases the cause of false-positive reactions is unclear (Avakian et al., 1988). In addition, cross-reacting antigens may be shared among mycoplasma species or between mycoplasmas and bacteria. Several cross-reacting antigens between M. gallisepticum and M. synoviae are known to occur. Different degrees of agglutination (intensity or speed of reaction) may be distinguished, allowing a semiquantitative scoring system. Some laboratories
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Stanley H. Kleven and Sharon Levisohn
determine agglutination end points by preparing double (1:2) dilutions of sera in saline. Sera that react at 1:8 or greater are considered positive, which also may be useful in differentiating between specific and nonspecific agglutination. Enzyme Linked Immunosorbent Assay (EUSA) Several laboratories have developed ELISA procedures; test kits for M. gallisepticum, M. synoviae, and M. meleagridis are also commercially available. Positive activity in the ELISA test develops shortly after SPA activity (about 2 weeks postinfection). Unfortunately, ELISA tests may also exhibit nonspecific positive reactions, although the commercially available kits are much more specific than earlier ELISA systems. ELISA tests based on the use of immunodominant species-specific proteins of M. gallisepticum or M. synoviae, or monoclonal antibodies to such proteins, have been developed and tested experimentally. Several additional tests based on the ELISA principle have been reported but have found limited use in field diagnosis. See Chapter B2 in this volume for further details on ELISA methodology. Hemagglutination-lnhibition Test Cultures of most strains of M. gallisepticum, M. synoviae, and some strains of M. meleagridis will agglutinate red blood cells of chickens, turkeys, and some mammals. Antibodies present in infected birds will inhibit the agglutination. The hemagglutination-inhibition (HI) test is more specific but less sensitive than the SPA and ELISA tests. Infected birds may not test positive until 3 weeks or longer after infection. In addition, there is antigenic variation among M. gallisepticum strains as measured by HI (Kleven et al., 1988). The HI test may be carried out with fresh culture of the hemagglutinating strain but most laboratories find it more convenient to use a standardized preserved antigen. Antigen may be prepared by harvesting an actively growing culture by centrifugation, suspending the packed cells in a small amount of phosphate-buffered saline, and mixing with an equal volume of glycerol. Aliquots are preserved by freezing at -70°C. Details of the HI test procedure using concentrated antigen can be found elsewhere (U.S. Department of Agriculture, 1993; Kleven and Yoder, 1989). Generally, HI titers of 1:40 to 1:80 or greater are considered positive, but the interpretation of results must be done on a flock basis. Tests for the presence of antibodies are usually carried out in serum, but yolk, blood, synovial fluid, and respiratory secretions may be used successfully in some tests. The presence of antibodies in these materials (specimens) has been demonstrated as well in other sites such as bile and Harderian glands, for which
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the indirect immunoperoxidase test may be used (Bencina and Bradbury, 1992). No serological tests for avian mycoplasmas other than M. gallisepticum, M. synoviae, and M. meleagridis exist.
Gene-Based Methods for Detection of Avian Mycoplasmas DNA probes have been designed for the detection of avian mycoplasmas, but the relative lack of sensitivity of such probes has inhibited their use for routine diagnostic purposes. PCR procedures for amplification of species-specific DNA sequences have been described for all the pathogenic avian mycoplasmas. Diagnostic methods based on direct demonstration of the specific DNA sequences have the advantage of being applicable under conditions where culture methods may not succeed, for instance, in mixed mycoplasma infections, high level of bacterial infection, or after antibiotic treatment of the flock. Nonviable mycoplasmas can also be detected as long as there has been no degradation of the DNA. However, not all of these advantages have yet been fully exploited since many of the currently available tests require a preliminary step of inoculation in culture medium before carrying out the test for the presence of the target gene. Nonetheless, these methods detect mycoplasma at early stages of infection (even before detection of antibodies) and in particular more quickly with than any other method. Commercially available kits (IDEXX, Westbrook, ME) are now being used routinely as a supplementary method for diagnosis while evaluation under field conditions continues. For a complete discussion of DNA probes and PCR, see Section A in this volume.
Summary Because of the economic importance of mycoplasmosis in the modem poultry industry, rapid and accurate diagnosis is essential. Although traditional serological testing methods have been useful in controlling infection in the past, there is emphasis on the development of improved diagnostic tools which are rapid, reliable, and economical. Together with this, research results have pointed to the complexity of diagnosis of the pathogenic species. Particularly withM. gallisepticum, marked intraspecies diversity has been found, at both the genetic and phenotypic levels. Developments in improving the understanding of the molecular biology of avian mycoplasmas will likely result in significant improvements in the ability to rapidly and economically detect infection in commercial poultry flocks. Because of the significance of vertical transmission via infected embryos
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and the need of maintaining poultry in large flocks, it should be emphasized that the diagnosis of mycoplasmosis must be made on a flock basis; the presence of one or more infected individuals in a flock indicates infection of the entire flock.
References Avakian, A. P., Kleven, S. H., and Glisson, J. R. (1988). Evaluation of the specificity and sensitivity of two commercial enzyme-linked immunosorbent assay kits, the serum plate agglutination test, and hemagglutination-inhibition test for antibodies formed in response to Mycoplasma gallisepticum. Avian Dis. 32, 262-272. Bendina, D., and Bradbury, J. M. (1992). Combination of immunofluorescence and immunoperoxidase techniques for serotyping mixtures of Mycoplasma species. J. Clin. Microbiol. 30, 407410. Bradbury, J. M. (1983). Phosphatase activity. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 363-366. Academic Press, New York. Clyde, W. A. Jr. (1983). Growth inhibition tests. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, 405-410. Academic Press, New York. Frey, M. L., Hanson, R. P., and Anderson, D. P. (1968). A medium for the isolation of avian mycoplasmas. Am. J. Vet. Res. 29, 2163-2171. Gardella, R. S., Del Giudice, R. A., and Tully, J. G. (1983). Immunofluorescence. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, pp. 431-439. Academic Press, New York. Jordan, F. T. W. (1983). Recovery and identification of avian mycoplasmas. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 69-79. Academic Press, New York. Jordan, F. T. W., and Amin, M. M. (1980). A survey of mycoplasma infections in domestic poultry. Res. Vet. Sci. 28, 96-100. Kleven, S. H., and Yoder, H. W. Jr. (1989). Mycoplasmosis. In "A Laboratory Manual for the Isolation and Identification of Avian Pathogens" (H. G. Purchase, L. H. Arp, C. H. Domermuth, and J. E. Pearson, eds.), pp. 57-62. Am. Assoc. Avian Pathol., Kendall/Hunt, Dubuque, lA. Kleven, S. H., Morrow, C. J., and Whithear, K. G. (1988). Comparison of Mycoplasma gallisepticum strains by hemagglutination-inhibition and restriction endonuclease analysis. Avian Dis. 32,731-741. Stipkovits, L., Varga, Z., Glavits, R., Ratz, F., and Molnar, E. (1987). Pathological and immunological studies on goose embryos and one-day-old goslings experimentally infected with a Mycoplasma strain of goose origin. Avian Pathol. 16, 453-468. U.S. Department of Agriculture (1993). "National Poultry Improvement Plan and Auxiliary Provisions," Publ. APHIS 91-40, pp. 54-63. Animal and Plant Health Inspection Service, U.S. Dept. of Agriculture, Hyattsville, MD.
D10 DIAGNOSIS OF SPIROPLASMA INFECTIONS IN PLANTS AND INSECTS C. Saillard^ C. Barthe, J. M. Bove, and R. F. Whitcomb
Introduction Diagnosis of spiroplasma infections in plants requires a thorough understanding of the infection process, epidemiology, and symptomatology of the pathogens in their respective hosts. At present, only three plant-pathogenic spiroplasmas are known. One of these, Spiroplasma phoeniceum, was recovered from natural and experimentally infected Catharanthus roseus, but has not been found in any other host (Saillard et al., 1987). The new spiroplasma was discovered and distinguished from S. citri by the fact that extracts from S. phoeniceuminfected periwinkle plants did not react in the ELISA with antiserum to S. citri. Likewise, extracts of S. c/rn-infected periwinkle plants gave negative reactions in ELISA tests with antiserum to S. phoeniceum. The com stunt spiroplasma {S. kunkelii) is frequently associated with other viruses and/or moUicutes in its major host Zea mays (Nault and Bradfute, 1979). In this case, the symptomatic picture in infected hosts may be truly confusing, and specific diagnostic tools are needed to make diagnoses with certainty (Whitcomb, 1989). ELISA techniques which employ polyclonal antibodies prepared from cultured organisms have proved to be effective in the diagnosis of S. kunkelii (reviewed by Whitcomb, 1989). The use of polyclonal antibodies, particularly those directed to group I organisms, raises the question of specificity. However, since no other spi293 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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roplasma is known to occur in maize in the wide range of com stunt in Latin America, ELISA gives unambiguous results. In contrast to S. kunkelii, S. citri occurs in a wide variety of hosts in nature. Epidemiological studies of this spiroplasma therefore require a detection method which is specific and which can be adapted readily to field-collected material. For many years, S. citri was detected in plants (citrus and periwinkles) and in insect vectors (leafhoppers) by culture of the organism on a selective growth medium (Bove et al., 1983). ELISA techniques (Saillard and Bove, 1983) were eventually developed for S. citri. However, neither culture tests nor ELISA permitted detection of S. citri in presymptomatic citrus plants. Failure to detect the agent in early stages of the plant disease is a result of the low number of spiroplasmas present in the phloem. A new test based on DNA probe technology has been developed that permits the detection of S. citri in infected plants prior to symptom development. This test utilizes the polymerase chain reaction (PCR), which increases the DNA copy number of a known sequence. From the nucleotide sequence of the spiralin gene (Chevalier et al, 1990) or S. citri virus SpVl DNA (Renaudin et al, 1990), suitable primers can be selected and used for PCR amplification. The use of PCR for the detection of S. citri in plants was developed on S. c/rr/-infected periwinkles. The sensitivity of this detection method was 100 to 1000 times higher than that of ELISA or culture assay. The PCR was performed on pellets of infected plant juices or diluted juices that had not been centrifuged. Amplified fragments were not obtained using healthy periwinkle juice or the juice of periwinkles affected by various phytoplasmas. Because crude plant extracts often inhibit PCR and decrease the sensitivity of the test, a new procedure, immunocapture PCR (IC-PCR), has been developed. This test simplifies sample preparation and enhances the specificity and sensitivity of conventional PCR. In ICPCR, spiroplasmas present in crude plant extracts are captured by polyclonal antibodies coating the walls of the tube in which the PCR is carried out. After the capture step, the plant extract is decanted and the spiroplasmas trapped in the tube are submitted to the PCR reaction without DNA isolation. Immunocapture of 5. citri cells by the coated antibodies and elimination of the plant extracts increases the sensitivity by a factor of 10. To assess the relative sensitivity of ICPCR, a comparative detection assay has been carried out among culture assay, ELISA, PCR, and IC-PCR using simulated infected samples (i.e., healthy citrus extracts to which known amounts of S. citri have been added). The lowest number of spiroplasmas that could be detected per milliliter of plant extract by ELISA tests was 10^-10^. In contrast, approximately 10"^ spiroplasmas per ml could be detected by PCR and 10^ by IC-PCR. This technique is therefore the most promising technique for the detection of low amounts of S. citri in citrus plants and insects.
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Materials Needles for collection of hemolymph. These are handmade from Pasteur pipettes. The tapered ends of the pipettes are heated and drawn to approximately one-tenth of their diameter in a cool gas flame. Finally, the tip of each needle is broken to give a very thin orifice. Plastic tubes, 0.5 ml Pipetman P200; P20 aerosol-resistant tips (Molecular Bioproducts, San Diego, CA) Petri dishes Razor blades Incubator, 37°C Thermocycler Eppendorf centrifuge Carbonate buffer: 2.93 g/liter NaHC03, and 1.53 g/liter NaC03, adjusted to pH9.6 Phosphate-buffered sahne (PBS) buffer; 8g/liter NaCl, 0.2 g/liter KH2PO4, 2.9 g/liter Na2HP04-12H20, and 2 g/liter KCl For PBS-sorbitol, add 70 g of sorbitol to 1 liter PBS PCR reaction mix (50-|xl reaction): 10 x reaction buffer (supplied by the manufacturer); Detergent Wl 1%, 2.5 |xl; Primers, 100 JJLM, 0.5 jxl each (selected according to Chevalier et al, 1990, or Renaudin et al., 1990); dNTP - mix 5 mM, 2.0 |JL1; Bovine serum albumin (BSA) (20 mg/ml) 1.0 jxl; dimthyl sulfoxide (DMSO), 2.5 |UL1; and water to 50 |xl. Reagents: Purified IgG, diluted in coating buffer (final concentration 6 |JLg/ml) Mineral oil Tag DNA polymerase (BRL)
Procedure Detection of S. citri in Plants by IC-PCR COATING OF PCR EPPENDORF TUBES (0.5 ml)
1. To each tube add 50 |xl of 6 |xg/ml purified anti-5. citri IgG in carbonate buffer. Leave at 37°C for 4 hours. 2. Remove IgG solution. Wash three times with 100 |xl of PBS-sorbitol buffer (10 minutes wash). Be certain that all liquid is removed after the final wash. At this stage the tubes can be kept frozen for several months at -20°C.
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PREPARATION OF PLANT EXTRACTS
1. With a razor blade, chop 1 g of sample (midribs, veins) to a fine mince in a sterile petri dish (diameter 6 cm) containing 1 ml of PBS-sorbitol buffer. 2. The mince can be left at room temperature for several hours. IMMUNOCAPTURE
1. With the tip of an Eppendorf pipette in contact with the bottom surface of the petri dish, remove 50 |xl of the minced tissue liquid and transfer to a precoated tube. Incubate overnight at 4°C. 2. Pour off the minced tissue fluid from the tube. Wash three times with 100 U | L1 of PBS-sorbitol buffer. Take care to decant the tube well. PCR REACTION
1. To each tube add 50 jxl of reaction mixture. Mix and spin down droplets in the Eppendorf centrifuge. 2. Overlay the reaction mixture with four drops of mineral oil to prevent evaporation. 3. Denature samples at 92°C for 30 minutes. Leave the tubes on ice for 10 minutes. Add 2.5 units of Taq DNA polymerase. 4. Perform 40 cycles of PCR using the following temperature profile. Denaturation: 92°C for 1 minute. Primer annealing: 52°C for 1 minute. Primer extension: 72°C for 2 minutes. Conclude cycles with a final extension at 72°C for 10 minutes. DETECTION OF 5. citri IN INSECTS BY PCR
Live adult leafhoppers are frozen for a few minutes at — 20°C. They are then placed dorsal side down on a glass slide. A needle is introduced into the abdomen; a volume of approximately 0.2 jxl is pipetted. These 0.2 |xl are placed in an Eppendorf tube (0.5 ml) and 50 |xl of the reaction mixture is added. The PCR reaction is carried out as described for plant samples. ANALYSIS OF PCR PRODUCTS
Ten microliters of the amplified DNA from the reaction mixture is analyzed on agarose or polyacrylamide gel according to standard procedures (Maniatis et al., 1982; see also Chapter A6, this volume).
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Discussion To date, PCR and IC-PCR are the most sensitive and specific techniques used for the detection of S. citri. PCR has the capacity to produce millions of DNA molecules. The main problems with the technique concern sample contamination from previous amplifications. Extreme care must be taken to avoid false-positive reactions. It is important to test the reagent controls for each amplification. One control must contain all PCR components except the template DNA. Furthermore, when PCR or IC-PCR is carried out on field trees it is obligatory to include healthy trees from the greenhouse as a control. If an amplified DNA fragment is visible on the gel from the healthy greenhouse trees, the presence or absence of S. citri in the field trees cannot be inferred. General procedures for minimizing the contamination in the PCR reaction have been described (Orrego, 1990).
References Bove, J. M., Whitcomb, R. F., and McCoy, R. E. (1983). Culture techniques for spiroplasmas from plants. In "Methods in Mycoplasmology" (J. G. TuUy and S. Razin, eds.), Vol. 2, pp. 225234. Academic Press, New York. Chevalier, C , Saillard, C , and Bove, J. M. (1990). Organization and nucleotide sequences of the Spiroplasma citri genes for ribosomal protein S2, elongation factor Ts, spiralin, phosphofructokinase, pyruvate kinase, and unidentified protein. J. Bacterial. 172, 2693-2703. Maniatis, T., Fritsch, E. F., and Sambrook, J. (1982). "Molecular Cloning: A Laboratory Manual." Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Nault, L. R., and Bradfute, O. E. (1979). Com stunt: Involvement of a complex of leafhopper borne pathogens. In "Leafhopper Vectors and Plant Disease Agents" (K. Maramorosch and K. F. Harris, eds.), pp. 561-586. Academic Press, New York. Orrego, C. (1990). Organizing a laboratory for PCR work. In "PCR Protocols" (M. A. Innis, D. H. Gelfand, J. J. Sninsky, and T. J. White, eds.), pp. 447-454. Academic Press, San Diego, CA. Renaudin, J., Aullo, P., Vignault, J. C , and Bove, J. M. (1990). Complete nucleotide sequence of the genome of Spiroplasma citri virus SPVL-R8A2B. Nucleic Acids Res. 18, 1293-1294. Saillard, C , and Bove, J. M. (1983). Application of ELISA to spiroplasma detection and classification. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 471-476. Academic Press, New York. Saillard, C , Vignault, J. C , Bove, J. M., Raie, A., Tully, J. G., Williamson, D. L., Fos, A., Gamier, M., Gadeau, A., Carle, P., and Whitcomb, R. F. (1987). Spiroplasmaphoeniceum sp. nov., a new plant-pathogenic species from Syria. Int. J. Syst. Bacteriol. 37, 106-115. Whitcomb, R. F. (1989). The biology of Spiroplasma kunkelii. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.), Vol. 5, pp. 487-544. Academic Press, San Diego, CA.
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D11 DETECTION OF PHYTOPLASMA INFECTIONS IN PLANTS E. Seemueller and B. C. Kirkpatrick
General introduction Methods for detecting plant-pathogenic phytoplasmas [formerly called mycoplasma-like organisms (MLOs)] in plants include microscopical, serological, and nucleic acid-based techniques. Transmission electron microscopy was the first method used for diagnosing phytoplasma infections in plant and insect hosts and it is still used today. Methods have also been developed to detect phytoplasma infections using bright-field and fluorescence microscopy. These methods, which have been reviewed by Chen et al., (1989), Hiruki (1988), and Waters (1982), include the observation of histochemical aberrations in infected tissue, as well as the direct observation of the organisms whose sizes are near the resolution limit of the light microscope. This chapter presents microscopic and nucleic acid-based methods that are now widely used to detect and characterize phytoplasmas in plants. For successful diagnosis of phytoplasma infections in plants one needs to keep in mind the phloem localization of the pathogens and the fact that pathogen titers can vary in different plant tissues. As phytoplasmas reside almost exclusively in sieve tubes, the samples should include as much phloem tissue as possible. This is particularly true for woody plants. Phytoplasma populations within the tree also vary during the growing season. In general, pathogen titers are highest in all portions of the tree in mid to late summer. The following papers provide specific details on phytoplasma colonization of herbaceous (Kuske and Kirkpatrick, 1992) and woody (Seemueller, 1988) plants. 299 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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DAPI Staining INTRODUCTION
Direct detection by light microscopy comprises the localization of aggregated phytoplasma cells with Dienes' stain (Deeley et al., 1979). This simple method, which has originally been developed to identify mycoplasma colonies on agar media, may still be useful for detecting phytoplasmas in high-titer plants; however, detection of phytoplasmas with DNA-binding fluorescent dyes has gained wider acceptance than bright-field methods in recent years. Of the various dyes that can be used, 4', 6-diamidino-2-phenylindole (DAPI) (Seemueller, 1976) became a standard in many laboratories and its use is described here. MATERIALS Fixative
Glutaraldehyde (EM grade) diluted to 5% (v/v) with 0.1 M phosphate buffer (38.57 g Na2HP04-12H20 and 12.46 g KH22PO4 per liter), pH 7.0. Alternatively, 4% (v/v) formaldehyde may be used. Staining Solution (100 ml)
1. Dissolve 0.1 mg DAPI in 100 ml phosphate buffer by stirring for 30 minutes at room temperature (thimerosal may be added to a final concentration of 0.01%). 2. Filter sterilize through a 0.22-|xm filter. Wrap the bottle in aluminum foil and store in the dark at 2° to 6°C as the stain is heat and light sensitive. Examine the solution periodically for bacterial contamination. Discard or filter sterilize if contaminated. Equipment
Freezing microtome Epifluorescence microscope equipped with a high-pressure mercury lamp (e.g., HBO50 or HBOIOO), an exciter filter transmitting between 300 and 400 nm (peak at ca. 365 nm; e.g., UGl), a barrier filter that transmits above 400 nm (e.g., K400 or K430), and two objectives [10 and 65x (60 to 80x)] recommended for UV application.
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PROCEDURE
1. Freshly collected roots, shoots, petioles, midribs, fruit peduncles, or phloem from the trunk or major limbs can all be examined by DAPI. Keep the samples turgid and cool and process them as soon as possible, especially phloem bark tissues. Wash roots thoroughly with water. 2. Cut specimens 5 to 8 mm in length and about 2 mm thick. Split shoots, petioles, peduncles, and finer roots longitudinally to enhance fixation. 3. Place samples immediately into vials with a generous amount of fixative. Fixative may be vacuum infiltrated to enhance fixation; however, fresh fixative should be added after vacuum infiltration. Store samples at 2° to 6°C. 4. Rinse specimens in 0.1 M phosphate buffer to remove the fixative and freeze it to the microtome stage with distilled water or a special tissue-embedding medium. Make radial sections, 15 to 25 |xm thick. 5. Transfer two to five sections to a microscope slide. Add DAPI solution to the slide so that all sections are covered. Stain for 2 to 5 minutes. Cover the sections with a coverslip using gentle pressure. Blot away any excess stain from the coverslip edges and seal the coverslip with nail polish. 6. For best results the sections should be examined immediately; however, they may be stored for up to 2 days in a tight vessel in the refrigerator. Use the 10X objective to find the current season's phloem, and use the high-power lens for the detection of phytoplasmas. DISCUSSION
DAPI binds specifically to DNA. However, like all DNA-staining procedures, DAPI is not specific for phytoplasmas. Although DAPI binds preferentially to (A+T)-rich DNA, DNA within plant organelles such as mitochondria and chloroplasts are also stained by DAPI. The specificity of DAPI examination for phytoplasma infection is obtained by preferentially examining the phloem tissues where the pathogen is localized. In longitudinal sections, sieve tubes can readily be recognized by their elongated shape and the presence of sieve areas in the radial cell walls and sieve plates. When completely mature, the sieve tube elements are largely free from cell organelles. Thus, there are essentially no fluorescent particles in mature sieve tubes of healthy plants. In contrast, small fluorescent particles can be observed in a variable number of sieve tubes in the current season's phloem of infected plants. In general, infection can be immediately recognized when phytoplasma numbers are sufficiently high; however, up to five sections from each of three randomly collected samples should be examined before a plant is considered to be negative. DAPI staining is a simple, rapid, inexpensive, and versatile method that
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permits pathogen detection in a sensitivity range that is similar to that obtained with nucleic acid hybridization procedures. It is suitable for the routine diagnosis of diseases of known etiology and for the screening of plant materials for the presence of phytoplasmas before additional time-consuming procedures such as DNA isolation or electron microscopy are carried out. However, to prove the phytoplasma etiology of a new disease, the DAPI results have to be confirmed by electron microscopy or molecular methods. Some modifications of the DAPI procedure have been reviewed by Chen et al., (1989) and Hiruki (1988). These methods included embedding of the tissue in paraffin, enzymatic maceration, mechanical crushing, and free-hand sectioning. However, all these modifications are less suitable than freeze sectioning because they are more time consuming, less sensitive, or the observations are more difficult to interpret. DAPI may be replaced by the benzimidazole Hoechst 33258, although it may cause more background fluorescence with plant tissue.
Detection of Phytoplasmas by DNA Hybridization Assays INTRODUCTION
The introduction of serological and DNA-based methods into phytoplasma diagnosis made it possible to detect specific phytoplasmas for the first time. Various serological methods such as the enzyme-linked immunosorbent assay (ELISA), immunofluorescence microscopy, immunosorbent electron microscopy, dot-blot immunoassay, and Western blot analyses are described in Chapter B6 and B8 of this volume. The methods of molecular genetics used in phytoplasma diagnosis include dot and Southern blot hybridization and polymerase chain reaction (PCR) technology. These procedures, which are described here, combine high sensitivity with a variable degree of specificity, depending on the sequences selected as probes or primers. In this way, conditions can be chosen for detecting most or all phytoplasmas, groups of phytoplasmas, or specific pathogens. Since the introduction of DNA technology into phytoplasma research in the late 1980s (Kirkpatrick et a/., 1987), DNA hybridization assays have become widely used to detect and characterize these organisms. Dot hybridization is a sensitive and relatively simple method which allows large numbers of samples to be screened quickly. A wide array of cloned chromosomal and extrachromosomal DNA probes have been developed which allow group-specific detection of phytoplasmas (Lee and Davis, 1988). Additional information is provided by Southern blot hybridization and analysis of resulting restriction fragment length polymorphisms (RFLP). Strains that were indistinguishable on the basis of dot hybridization assays could be differentiated on the basis RFLP analysis. There-
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fore, RFLP analysis has become a suitable means for identifying specific phytoplasma strains. No standard protocols exist for performing DNA hybridization analyses. Choices must be made between different types of hybridization and transfer buffers, membranes, DNA immobilization methods, and probe-labeling protocols. Although nitrocellulose and uncharged nylon membranes are interchangeable for most applications, modifications are required when positively charged nylon membranes are used. Oligonucleotide and RNA probes as well as chromosomal and extrachromosomal DNA probes have been described. However, for most routine detection studies we prefer randomly cloned fragments of the phytoplasma chromosome that are between 1 and 5 kb. Labeling may be carried out either radioactively or nonradioactively, but most nonradioactive detection systems are less sensitive than ^^p autoradiography. However, nonradioactive hybridization probes are safer to prepare and to handle and can be stored for years. Of the nonradioactive systems, the chemiluminescent detection is considerably more sensitive than the colorimetric detection and it has the advantage that reprobing the membranes is possible. The following protocols use uncharged nylon membranes. Southern transfer of DNA fragments from the agarose gel to the membrane is accomplished using the traditional upward capillary transfer method. DNA can be bound to the membrane by heat or UV irradiation, and the technique uses a radioactively labeled DNA hybridization probe. Information on modifications required by the use of other membranes as well as transfer and immobilization methods is mentioned. For details on other labeling procedures and the use of oligonucleotide and RNA probes, the reader is referred to Chapters A2 and A3 in this volume and to the manuals by Sambrook et al (1989) and Ausubel et al. (1987, 1993). Dot'Blot Analysis of Phytoplasma DNA MATERIALS
Sample DNA (see Chapter B2 in Vol. I, for preparation) 20X SSC (IX SSC: 0.15 M NaCl, 0.015 M sodium citrate, pH 7.0) 2 M Tris, pH 7.0 Uncharged nylon (or nitrocellulose) membrane Dot-blot or slot-blot manifold UV transparent plastic wrap (e.g., Saran wrap) UV transilluminator or UV light box Boiling water bath or heating block Standard automatic pipette and tips Microcentrifuge and sterile disposable tubes
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PROCEDURE
1. Soak the nitrocellulose or nylon membrane in 8x SSC for at least 15 minutes before applying the DNA samples. Put the membrane into the dot/slotblot apparatus following the manufacturer's protocol. 2. See Chapter B2 in Vol. I for DNA preparation. Suspend the DNA sample in 8x SSC or add 20 x SSC to the sample to give a final concentration of 8x SSC. 3. Denature the DNA by boiling for 5 minutes in the presence of 3 |JL1 of 2 M NaOH per 50 |xl of sample. Place in ice and neutralize by adding 3 |JL1 (per 50 |JL1) of 2 M Tris-HCl, pH 7.0. 4. Spin the samples (1 to 5 |xg of nucleic acids in the undiluted samples) in a microfuge for 5 seconds to sediment small particles which may block the membrane. If desired, make twofold serial dilutions of the sample in 8x SSC. Carefully apply the samples to the membrane without touching the membrane. 5. After all of the samples have been applied to the membrane, dissemble the dot/slot-blot apparatus and let the membrane dry at room temperature. 6. Immobilize nucleic acids by baking under vacuum on a gel dryer at 80°C for 30 minutes. Alternatively, the DNA may be fixed to the membrane by exposing the membrane to UV irradiation. Wrap the membrane in UVtransparent plastic wrap, place on a UV transilluminator (sample side down), and irradiate for 30 seconds to 5 minutes, depending on the power of the machine. Store the membrane at room temperature in a desiccator.
Southern Blot Analysis of Phytoplasma DNA MATERIALS
Sample DNA (see Chapter B2, Vol. I, for preparation) Restriction endonucleases such as EcoRl or Hindlll which efficiently cut A:Trich DNA 10 X restriction endonuclease buffer (supplied by enzyme manufacturer) lOx gel-loading buffer (20% Ficoll 400, 0.1 M EDTA, pH 8.0, 0.25% bromphenol blue) IX TAB or IX TBE electrophoresis buffer ( i x TAE: 40 mM Tris-acetate, 1 mM EDTA, pH 8.0; i x TBE: 90 mM Tris-borate, 90 mM boric acid, 2 mM EDTA, pH 8.0) Ethidium bromide (stock solution 10 mg/ml) Electrophoresis-grade agarose DNA molecular weight markers 0.25 M HCl
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Denaturation solution (1.5 M NaCl/0.5 M NaOH) Neutralization solution (1.5 M NaCl/0.5 M Tris-Cl, pH 7.0) 20X SSC (IX SSC: 0.15 M NaCl/0.015 M sodium citrate, pH 7.0) Whatman 3MM filter paper sheets Nylon (or nitrocellulose) membrane UV-transparent plastic wrap (e.g., Saran wrap) Horizontal gel electrophoresis apparatus Power supply UV transilluminator Polaroid camera equipped with Kodak Wratten No. 23A and No. 2B filter Polaroid type 667 film PROCEDURE
1. Pipette 2 |xl of lOx restriction buffer, 1-10 |xg of sample DNA, and 5 U of restriction endonuclease per microgram of DNA in a microfuge tube. Add H2O to a final volume of 20 jxl and incubate at the recommended temperature (usually 37°C) for 4 hours to overnight. Stop reaction by adding 2 jxl of 10 x gel-loading buffer. If the sample DNA is to be digested with multiple enzymes, digestions may be performed simultaneously if the buffers for the restriction enzymes are compatible. If the enzyme buffers are too dissimilar, digest successively by starting with the enzyme that requires the lowest concentration of salt, then add additional NaCl and the second enzyme. 2. Prepare a 0.7 to 1.0% agarose gel using 1 x TAE or TBE buffer. Separate the DNA fragments generated by restriction digestion by electrophoresing them through the gel using the same buffer. Electrophorese the sample until the bromophenol dye is approximately 1 cm from the bottom of the gel. 3. Stain the DNA fragments by adding ethidium bromide to a final concentration of 0.5 |xg/ml in the gel (do this when preparing the gel) or by soaking the gel after electrophoresis in an ethidium bromide solution (0.5 |xg/ml) for 10 minutes. Rinse the gel briefly with distilled water. Visualize the stained fragments using a UV transilluminator and photograph the gel with a Polaroid camera using Type 665 or 667 film. 4. If desired, partially depurinate the DNA by soaking the gel in several volumes of 0.25 M HCl for 10 to 15 minutes (this step can be omitted when efficient transfer of molecules >10 kb is not required). Rinse the gel in distilled water and soak it in several volumes of denaturation solution for 45 minutes. Then rinse the gel in distilled water and soak it in several volumes of neutralization solution for 30 minutes. Replace the neutralization solution and continue soaking for a further 20 minutes. Perform depurination, denaturation, and neutralization with gentle agitation at room temperature. 5. Transfer the denatured DNA fragments to a nylon or nitrocellulose mem-
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brane using the upward capillary transfer method (Sambrook et al, 1989) for a minimum of 12 hours. Use 8x SSC as the transfer buffer for nitrocellulose or uncharged nylon membranes and 0.4 M NaOH for positively charged nylon membrane. When transfer is complete, briefly rinse the membrane in 2x SSC to remove residual agarose. Dry the membrane at room temperature and bind the DNA by baking or UV cross-linking as previously described for dot-blots.
Hybridization Analysis of Dot or Southern Blots MATERIALS
Membrane with dot or Southern blot samples (see earlier) Cloned phytoplasma DNA probe (see Chapter A3, this volume) [a-32p]dATP Nick translation or random priming labeling kit 20 X SSC (see earlier) lOOx Denhardt's solution [2% (w/v) Ficoll 400, 2% (w/v) polyvinylpyrrolidone (molecular weight 10,000), 2% (w/v) bovine serum albumin (fraction V)] 10% solution of sodium dodecyl sulfate (SDS) in H2O Salmon or herring sperm DNA Scintillation counter Hybridization oven or 68°C water bath or incubator Hybridization tubes or scalable bags and heat sealer Film cassette for X-ray film with intensifying screen (e.g., DuPont Cronex Lightening Plus or Fuji Mach 2) X-ray film (e.g., Kodak X-OMAT R or Fuji RX) PROCEDURE
1. Isolate the probe DNA from recombinant plasmids by digesting the recombinant plasmid with enzymes that will cut out the insert. Electrophorese the digest in low-melting or standard agarose gels and recover the cloned insert by cutting out the cloned DNA fragment from the gel and removing the agarose matrix using the GeneClean System (Bio 101, La JoUa, CA), electroelution, or agarose-digesting enzymes such as gelase (EpiCentre Technologies). Alternatively, the cloned probe can be generated by PCR amplification (as described in Chapters A3 and A4, this volume). 2. Label the DNA to a specific activity of 10^ to 10^ dpm/jxg by nick translation or random oligonucleotide priming according to the instructions provided in the labeling kit.
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3. Place the membrane in a hybridization tube or a polyethylene bag. Add prehybridization solution (1 ml per 10 cm^ of membrane) consisting of 6x SSC, 5x Denhardt's solution, 0.5% SDS, and 100 |xg/ml of denaturated salmon sperm DNA and incubate for 3 hours with rotation (or gentle shaking) at 55°C. 4. Replace the prehybridization solution with an equal volume of a prewarmed (55°C) solution of the same composition. Just before use, denature the double-stranded probe by boiling for 10 minutes. Immediately add the denaturated probe to the hybridization solution and incubate overnight (16 hours) at 55°C. 5. Replace the hybridization solution with an equal volume of 2x SSC/0.1% SDS and incubate with rotation for 15 minutes at room temperature. Change the solution and repeat washing under the same conditions. Take care to avoid letting the membrane dry when changing the wash solutions. 6. Carry out a further two washes as described in step 5, using prewarmed 0.2X SSC/0.1% SDS for 30 minutes at 55°C (moderately stringent conditions). If desired, further washes at a higher stringency may be carried out with prewarmed 0.1X SSC/0.1% SDS for 30 minutes at 60°, 65°, and/or 68°C (twice at each temperature). 7. After the final wash, rinse the membrane in 2x SSC at room temperature. Blot excess liquid and wrap the membrane in plastic wrap. Place the wrapped membrane in a cassette and expose it to X-ray film using an intensifying screen at -70°C. 8. If desired, the membrane can be rehybridized after removing the bound probe. This is accomplished by pouring several hundred milliliters of hot (90°C) solution containing 1% SDS/O.Olx SSC onto the membrane. Monitor the removal of the probe with a hand-held Geiger counter and repeat as necessary. The membrane must not be allowed to dry out between hybridization and stripping. Expose the stripped membrane to X-ray film to verify complete removal of the probe. Reinforced nitrocellulose or nylon membranes are preferred for rehybridization because they withstand the extreme temperatures better than standard nitrocellulose. DISCUSSION
The detection and identification of phytoplasma infections in plants by hybridization assays may encounter several problems. One of them is sensitivity. The amount of phytoplasma DNA in total DNA from infected plants is low, especially in many woody hosts. Usually, no hybridization signals are obtained when phytoplasmas cannot be detected by DAPI staining. When the phytoplasma concentration is low, MLO-enrichment procedures such as those described by Kirkpatrick et al. (1987) and Ahrens and Seemueller (1992) (see Chapter B2, Vol. I) should be used. DNA preparations from plant hosts may contain impuri-
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ties that affect dot hybridization results. These impurities may reduce hybridization signals by inhibiting probe hybridization or, alternatively, they may nonspecifically stick to the probe, thus producing false-positive signals. These problems can be circumvented by electrophoresing undigested sample DNAs in an agarose gel and blotting the DNA onto a membrane. Impurities from plants may also inhibit restriction endonuclease activity. The decreased activity may be compensated for by increasing the amount of enzyme added to the reaction mixture (up to 10-20 U/|xg DNA), increasing the reaction volume to dilute inhibitors, or increasing the duration of incubation. Also, the digestion of DNA can be facilitated by adding the polycation spermidine (final concentration 1 to 10 mM) which binds negatively charged contaminants. Other methods for removing contaminants from plant DNA include gel filtration through a Sepharose CL-4B (Parmacia) column or purification by ion-exchange resins such as an Elutip-d column (Schleicher and Schuell).
Analysis of Phytoplasmas Using Polymerase Chain Reaction INTRODUCTION
PCR technology offers several advantages for phytoplasma detection and characterization, including high sensitivity, versatility, relative simplicity, and nonradioactive procedures. PCR can reliably detect low-titer phytoplasma infections in plants that cannot be detected using microscopic, serological, or hybridization methods. Basic information on the application of PCR technology in mollicute research is given in Chapters A4 to A8 in this volume. Although the selection of target sequences is the subject of one of these chapters (A4), some of those aspects relevant for the detection of phytoplasmas will be discussed here. PCR primers obtained from phytoplasma 16S rRNA gene sequences are the most extensively used. 16S rDNA has highly conserved, semiconserved, and variable regions which offer the possibility for designing primers that have a wide or narrow range of specificity, thus amplifying the corresponding sequences of most or all phytoplasmas, or only a certain pathogen or a narrow group of pathogens. Primers with usually higher specificity were designed from sequences of the spacer region between the 16S and 23S rRNA genes (Kirkpatrick et al., 1994) because this region is more variable than the 16S rRNA. Highly specific and sensitive detection was also achieved using primers whose sequence was obtained from randomly cloned phytoplasma DNA fragments. The following protocol is a general one that uses primers obtained from the 16S rRNA gene. The identity of the specific primer sequence is presented in Chapter E6 of Vol. I, which discusses phytoplasma phylogeny and taxonomy.
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MATERIALS
Template DNA from healthy and phytoplasma-infected plants (see Chapter B2, Vol. I) Sterile H2O 10 X amplification buffer 2 mM dNTP mix [2 mM of each dNTP in TE buffer (10 mM Tris, 1 mM EDTA, pH 7.5)] 10 |xM of each oligonucleotide primer Taq DNA polymerase Light mineral oil Automated thermal cycler Sieving agarose (e.g., FMC Metaphor or NuSieve 3:1 agarose) Reagents and equipment for agarose gel electrophoresis as described under Southern blotting PROCEDURE
1. In a sterile microfuge tube mix 20 jxl of sterile H2O, 4 |xl of lOx buffer, 2 |JL1 of dNTP mix (final concentration 100 jxM), 2 |JL1 of each primer (final concentration 0.5 JJLM), about 50 to 200 ng of template DNA (depending on the relative concentration of phytoplasma DNA in the extract), 1 U Taq polymerase, and bring to a final volume of 40 \x\ with sterile H2O. Set up two negative controls, one with sterile water instead of template DNA and one with DNA from a corresponding healthy plant. Include a positive control containing template DNA from a known infected plant that contains the target sequence. Overlay the reaction mixture with 40 JJLI mineral oil. 2. Carry out 30 to 35 amplification cycles at the following incubations: 1 minute of denaturation at 94°C (5 minutes for the first cycle), 1 minute of primer annealing between 50° and 60°C depending on the T^ of the primers, and 1 to 3 minutes of strand extension at 72°C (approximately 1 minute/kb of amplified product). The extension time of the last cycle should be increased to 7 minutes to make products as complete as possible. 3. Analyze the PCR products by electrophoresing 10 jxl of the reaction mixture in a 1% agarose gel; visualize the products by staining the gel with ethidium bromide and UV illumination as previously described. Use sieving agarose gel for resolving small PCR products (<500 bp). Alternatively, small PCR products can be resolved by PAGE (see Chapter A6, this volume). 4. The amplification products may be characterized further and the phytoplasma taxonomically classified by sequencing or digesting the PCR product with restriction endonucleases such as Alu\ or Rsal (Schneider et al., 1993). Specific details on sequencing or analyzing PCR-amplified phytoplasma 16S rRNA genes are presented in Chapter A6 of this volume.
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DISCUSSION
PCR is an extremely sensitive method for detecting phytoplasma; however, contaminants in the DNA preparation can severely inhibit the amplification reaction. Occasionally, the efficiency of amplification can be improved by diluting or preboiling the sample or by using different extraction DNA procedures. Inhibitors can often be removed by Elutip-d (Schleicher and Schuell, Keane, NH) column purification or by centrifugation through Sephadex G-200 columns (Pharmacia, Piscataway, NJ). Increasing the number of amplification cycles (up to 40) can add sensitivity; however, the risk of amplifying contaminating DNA is increased as well. The specificity of detection can be improved by avoiding or reducing mispriming. This is achieved by adding the Taq polymerase to the reaction mixture before or after the first denaturation step at temperatures between 60° and 94°C ("hot start"). Mispriming can also be reduced by raising the annealing temperature during the first 10 cycles by 4° to 7°C above the "normal" annealing temperature. Furthermore, a double amplification, first with peripheral and then with internal primers, improves the specificity (nested PCR; Chapter A7, this volume).
References Ahrens, U., and Seemueller, E. (1992). Detection of DNA of plant pathogenic mycoplasmalike organisms by a polymerase chain reaction that amplifies a sequence of the 16S rRNA gene. Phytopathology 82, 828-832. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K., eds. (1987). "Current Protocols in Molecular Biology." Current Protocols, Brooklyn, NY. Ausubel, F. M., Brent, R., Kingston, R. E., Moore, D. D., Seidman, J. G., Smith, J. A., and Struhl, K., eds. (1993). "Current Protocols in Molecular Biology." Current Protocols, Brooklyn, NY. Chen, T. A., Lei, J. D., and Lin, C. P. (1989). Detection and identification of plant and insect mollicutes. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.). Vol. 5, pp. 393424. Academic Press, San Diego, CA. Deeley, J., Stevens, W. A., and Fox, R. T. V. (1979). Use of Dienes' stain to detect diseases induced by mycoplasmalike organisms. Phytopathology 69, 1169-1171. Hiruki, C. (1988). Fluorescence microscopy of yellows diseases associated with plant mycoplasmalike organisms. In "Mycoplasma Diseases of Crops" (K. Maramorosch and S. P. Raychaudhuri, eds.), pp. 51-76. Springer-Verlag, New York. Kirkpatrick, B. C , Smart, C. D., Blomquist, C , Guerra, L., Harrison, N. A., Ahrens, U., Lorenz, K.-H., Schneider, B., and Seemuller, E. (1994). Identification of MLO strain-specific PCR primers obtained from 16/23S rRNA spacer sequences. lOM Letters 3, 261-262. Kirkpatrick, B. C , Stenger, D. C , Morris, T. J., and Purcell, A. H. (1987). Cloning and detection of DNA from a nonculturable plant pathogenic mycoplasma-like organism. Science 238, 197200.
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Kuske, C. R., and Kirkpatrick, B. C. (1992). Distribution and multiplication of western aster yellows MLO in Catharanthus roseus determined by DNA hybridization analysis. Phytopathology 82, 457-462. Lee, I.-M., and Davis, R. E. (1988). Detection and investigation of genetic relatedness among aster yellows and other mycoplasmalike organisms by using cloned DNA and RNA probes. Mol. Plant-Microbe Interact. 1, 303-310. Sambrook, J., Fritsch, E. P., and Maniatis, T., eds. (1989). "Molecular Cloning: A Laboratory Manual," 2nd ed. Cold Spring Harbor Laboratory, Cold Spring Harbor, NY. Schneider, B., Ahrens, U., Kirkpatrick, B. C , and Seemueller, E. (1993). Classification of plantpathogenic mycoplasmalike organisms using restriction site analysis of PCR-amplified 16S rDNA. J. Gen, Microbiol 139, 519-527. Seemueller, E. (1976). Investigations to demonstrate mycoplasmalike organisms in diseased plants by fluorescence microscopy. Acta Hortic. 67, 109-111. Seemueller, E. (1988). Colonization patterns of mycoplasma-like organisms in trees affected by apple proliferation and pear decline. In "Tree Mycoplasmas and Mycoplasma Diseases" (C. Hiruki, ed.), pp. 179-192. Univ. Alberta Press, Edmonton, Alberta. Waters, H. (1982). Light and electron microscopy. In "Plant and Insect Mycoplasma Techniques" (M. J. Daniels and P. G. Markham, eds.), pp. 101-151. Croom Helm, London.
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D12 IDENTIFICATION OF MOLLICUTES FROM INSECTS Robert F. Whitcomb and Kevin j . Hackett
Introduction After the initial discovery that spiroplasmas were associated principally with arthropods, major efforts were made to isolate mollicutes from diverse insect taxa and from ecosystems that the insects inhabited (Hackett and Clark, 1989). These studies resulted in the discovery of a large number of Spiroplasma species and several additional genera of mollicutes. One of these genera (Acholeplasma) appears to inhabit various niches in nature, including vertebrate animals, insects, and plant surfaces. In rare cases, members of the genus Mycoplasma have apparently been isolated from plants and insects, as have some L-phase variants of bacteria. The latter two types of wall-less prokaryotes are probably not associated with arthropods, but have been encountered as artifacts of the isolation procedure. Other taxa have proved to be predictably associated with arthropods, especially insects. These taxa have formed the basis for a new order of arthropod-associated mollicutes, the Entomoplasmatales (Tully et al., 1993). Techniques for isolation of mollicutes from insects (Markham et al., 1983; Hackett and Clark, 1989) and for preparation of media suitable for isolations from these habitats are reviewed in Volume I of this series (see Chapters A2, A3, and A4) and in chapter DIO of this volume. This chapter describes the identification procedures for wall-less prokaryotes isolated from arthropods.
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Steps to Be Taken before Identification Begins If only a few isolates are obtained in a given study, it is likely that each will be important enough to merit detailed study. In this case, each isolate should be cloned (see Chapter E2 in Vol. I). Motile, helical organisms that are absolutely resistant to penicillin can be tentatively presumed to be spiroplasmas. If they turn out to represent a new group or subgroup, the lack of cell wall must be confirmed by electron microscopy (see Chapter E2 in Vol. I) in the course of description of the new group (Whitcomb et al., 1987). If the organisms are nonhelical, some consideration should be given to the possibility that they could be bacterial L-phase variants (see chapter E2, Vol. I). If this is deemed unlikely, the organisms should be passed to a bank of diagnostic media, including serum-containing and serum-free media with or without a polysorbitan monooleate (Tween 80) supplement (see Chapter E7, Vol. I). From these preliminary tests, the genus can be tentatively assigned. In some studies, the isolation rate may be so high that it is not feasible to clone every isolate. If this is true, the possibility that diagnostic tests have been performed on a mixed culture must be taken into account. In the section on identification by the spiroplasma deformation test (see the following section), the detection of mixtures is discussed.
Identification of Genera of Wall-less Mollicutes from Insects Identification of mollicutes from insects has become increasingly complex; new genera of organisms have been discovered in arthropods and species and groups from the diverse genera have proliferated. The following key gives a guide to the principles involved in distinguishing the genera of mollicutes associated with arthropods. Key to Genera of Wall-less Prokaryotes from Arthropods 1. 2.
3. 4.
Helical Spiroplasma Nonhelical 2 Passes 450-(xm but not 220-|jLm filter; reverts to walled bacterium when passed in conventional antibiotic-free bacteriological media Bacterial L-phase variant Filterable through 220-|xm filters; penicillin-resistant; verified by electron microscopy to be wall-less 3 Grows in serum-free mycoplasma broth Acholeplasma Does not grow in serum-free broth 4 Grows in serum-free broth supplemented with 0.04% Tween 80 Mesoplasma Does not grow in serum-free broth with Tween 80 5
D12 Identifying Mollicutes from Insects 5.
6.
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Reverts to helical morphology at some stage in growth cycle when tested with various media Spiroplasma Does not revert 6 Grows optimally at 37°C, possible contaminant from serum in medium or from the ecosystem Mycoplasma Grows optimally at a temperature less than 37°C, often 30° or 32°C; may grow at 20°C or less Entomoplasma
Identification to Species or Group Level Although the species is the central unit in mollicute taxonomy, it is an arbitrarily defined unit. Several features characterize the species. The first is that the clusters of strains that constitute mollicute species differ from other clusters by at least 30% of their genome (i.e., there is less than 70% DNA-DNA homology as measured by hybridization studies). Description of new species must conform to minimal standards (see Chapter E2 in Vol. I). The species (or, in the case of spiroplasmas, the group) is determined, as a practical matter, by serology. When the genus of a strain has been determined, the culture should be tested against specific antisera directed to the known species of that genus. Nonhelical organisms should be tested by the growth inhibition test (Tully et aL, 1994). Helical organisms should be tested by the spiroplasma deformation test, as detailed later. Antisera are available, at present, to 12 Mesoplasma, 5 Entomoplasma, and 13 Acholeplasma species. In the unlikely event that an isolate should turn out to be a true Mycoplasma species, it must be tested against sera to about 100 species. If the organism is helical, sera are available for 25 groups (17 of which have been given binomial names), 11 unique subgroups, and 8 putative groups that are well characterized (Tully et aL, 1987; Whitcomb et aL, 1992b; See also Appendix, this volume). In tests with particular Spiroplasma or Mycoplasma species, it is sometimes important to use antiserum to one or two other strains within the species complex. This ensures that the test spans the antigenic variability of the species; failure to do so could result in missing the identity of the unknown organism. Species and strain designations of mollicutes are given in the Appendix to this volume.
Identification of Infraspecific Units Various hierarchies of variation have emerged from the detailed study of mollicute strains (Whitcomb, 1994). In spiroplasmas, the group is a cluster of strains that is serologically unrelated to all other groups. In cases where DNA-
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DNA homology data are available, groups have turned out to share little or no homology; thus the spiroplasma serogroups can be thought of, tentatively, as homology groups. The term subgroup has been used to denote clusters of strains that share a considerable amount of DNA-DNA homology, but that are nonetheless substantially different (Whitcomb et al., 1987). The subspecies designation in mollicutes has been reserved for special cases in which it has been desirable to construct a formal taxonomy for economically important clusters of strains that share more than 70% DNA-DNA homology, but less than about 80%. In many spiroplasma assemblages (e.g., those from horseflies and mosquitoes, and the various representatives of group IV spiroplasmas from a number of sources), a wide range of variability has been encountered. Variability in these groups, although less than that between subgroups, is still substantial and appears to reflect different habitats and ecologies from which the variants have been derived. Such variation can be described at the serovar level. Spiroplasma serovars described by deformation tests can be described in terms of their reactivity with several antisera (see the next section). In many cases, there will be a lack of background information on the meaning of partial serological reactions. If such is the case, identifications will of necessity be only partial; many isolates may be identified as being serologically related to the representative strain of a known group, but not identical to it. A discussion of the issues involved in infraspecific variation is beyond the scope of this chapter, but is presented in several reports (Whitcomb et al, 1987; Whitcomb, 1994).
Spiroplasma Deformation Test Adaptation of the spiroplasma deformation test to routine diagnosis has resulted in significant modifications to previously reported protocols (Williamson and Whitcomb, 1983). We present the protocol for the test as currently performed. Materials Culture of an unknown spiroplasma in logarithmic growth phase in MID medium (Whitcomb, 1983); some tick-associated spiroplasmas may require SP-4 medium (see Chapter A2 in Vol. I) Dilutions of antiserum specific to each group (25 sera), subgroup (11 sera), and putative group (8 sera), in twofold dilutions (1:10 to 1:20,480) in MID medium Polyvalent antisera designed in accordance with past cultural experience 96-well microtiter plates
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Automatic pipetter Dark-field microscope
Procedure ANTIGEN-ANTIBODY REACTION
1. Pipette 25 jxl of each of the 12 antiserum dilutions into a single row of wells in the microtiter plates. 2. Adjust antiserum dilutions to about 10^ helical cells/ml (Rodwell and Whitcomb, 1983). 3. Add 25 fxl antigen to each antiserum dilution. 4. After 30 minutes, examine antigen-antibody mixtures under dark-field microscope. CHOICE OF TEST SERA
In many cases, a background of previous data produces an expectation of isolation for certain spiroplasma groups and/or species. When this is true, it is possible to screen isolates against a reduced bank of antisera and to further reduce the work volume by use of polyvalent sera constructed by mixing univalent, independently produced antisera. For example, six spiroplasma groups are regularly isolated from horseflies in the southeastern United States (French et al., 1990; Whitcomb et al., 1992a). We screen new isolates against a bank of only five polyvalent sera. One of these contains antibodies at a final dilution of 1:10 to the B31T (S. apis), W13, and PPSl strains of group IV; although this group appears in tabanids in northern latitudes and in Europe (Le Goff et al., 1991), it is very rare in tabanids in the southeastern United States. The second polyvalent antiserum contains antisera to the three representative strains of the subgroups of spiroplasma group VIII: strains EA-1 (VIII-1), DF-1 (VIII-2), and TAAS-1 (VIII-3) (Gasparich et al., 1993). The third polyvalent serum contains sera directed against the group XIV EC-1 strain and the ungrouped HYOS-1 strain. The fourth polyvalent serum contains antibodies directed against the group XXIII strain TG-1 and the ungrouped TAUS-1 strain. Most strains from the southeast react with one of the last three polyvalent sera. Occasionally, new isolates react with polyvalent serum number 5, which contains a mixture of antibodies, each a dilution of 1:10, against the group XVIII TN-1 strain and ungrouped strains TALS-2 and TABS-2; each of the latter two strains is a candidate for status as a new spiroplasma group. This typing scheme is adequate for tabanid spiroplasma assemblages from the southeastern United States. In a different geographical region, the experience may be quite different. Strains
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^!/,f'A!
Fig. 1. Diagrammatic representation of various microscopic observations obtained in spiroplasma deformation tests carried out under dark-field microscopy. (A) In the absence of a specific antibody, the culture consists predominantly of helical, motile cells; (B) end point of a deformation test, at which approximately one-half of the cells are deformed and exhibit blebs; (C) high levels of antibody induce immobilization and multiple blebbing, vi'ith agglutination of affected cells; and (D) evidence of mixed culture where the field contains some motile and helical unaffected organisms, but also aggregated and multiply blebbed cells.
isolated from horseflies in Europe exhibit a very different profile (Le Goff et al., 1991; C. Chastel, personal communication). READING THE TEST
1. If the culture contains largely typical, helical motile spiroplasma cells (Fig. lA), the test can be read as previously described (Williamson and Whitcomb, 1983). Briefly, the dilution at which about one-half of the helical cells have been deformed (Fig. IB) is recorded, and the titer of the reaction is expressed as the reciprocal of the end-point dilution.
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il^
Fig. 1. {Continued)
2. In many primary cultures, all or many spiroplasma cells are not perfect helices. It is common for many cells to be filamentous, to have a certain amount of nonspecific blebbing, or to have many very short or almost completely nonhelical forms. In all of these cases, it is possible to recognize serological deformation, providing a control culture that has not reacted with a specific antibody is available. In most cases, the well with the lowest antibody-antigen ratio (i.e., well No. 12, total dilution 1:41,960) will serve as this control. In a few cases, in which serological reactions may occur at very dilute antiserum concentrations, it may be necessary to use a control consisting of cultured organisms to which an equal volume of medium has been added. Ideally, in this control, there will be a large population of unaffected cells (Fig. 1 A). If more than one set of antibodies is utilized, the negative reactions from antisera that do not react with the unknown spiroplasma serve as auxilliary controls; these reactions control, to some extent, for the possible effect of a nonhomologous antibody and other components that may be present in heterologous antisera.
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3. In reading the results, it is important to be thoroughly familiar with the various types of spiroplasma-antibody reactions. At very high levels of antibody attack, some spiroplasmas (e.g., S. clarkii) are immobilized without blebbing; it would be possible to mistake this response for a negative reaction. At high levels of antibody attack, the aspect of the affected cells is one of multiply blebbed cells, with varying degrees of agglutination between affected cells (Fig. IC). As a result, the total number of bodies in the reaction mixture is greatly reduced, and no free helical filament, attached or unattached to any bleb or bleb mass, is found in the culture. As the ratio of antibody with respect to antigen decreases, a point is reached at which some cells have portions of helical filaments that are unaffected by antibody. This is true, even though 100% of the cells may show some effect of antibody. At somewhat lower levels of antibody, some cells escape attack altogether; eventually, the fraction of affected cells decreases to about half—this is the end point. Affected cells, although constituting less than 50% of the population, are usually evident at antibody attack rates one-fourth or, rarely, one-eighth those at the end point. 4. Although mixed infections are always of some concern in series of primary isolations, certain hosts carry frequent infections of different spiroplasmas. Such cases offer serious challenges in serotyping of primary isolates. In other mollicute systems, in which primary isolations are carried out on solid media, mixtures can be readily detected by use of fluorescent antibody techniques (see Chapter B9, this volume). Because spiroplasmas produce colonies erratically if at all, few if any spiroplasmas are isolated on solid media. The problem is, then, to recognize a mixture during primary isolation in liquid media. This has been accomplished in our laboratory by noting the morphology of affected spiroplasmas. The reaction series of an unmixed isolate, or of a culture that has a great predominance of a single spiroplasma serovar, is usually seen as a progression, from aggregated multiply blebbed cells, to populations of cells, some of which have single blebs. The identity of such cultures can be tentatively established by normal deformation tests. In contrast, the presence in a single antigen-antibody reaction of aggregated multiply blebbed cells and free, unaffected cells (Fig. ID) is a clear warning of a mixed culture. If such a culture is passed 5-10 times, the serological profile usually shifts, as one of the components of the original mixture eventually outgrows the other. Thus, a shift in apparent serological identity during cultural history is a second warning of a mixed culture. Resolution of such a mixture to identify all spiroplasmas may be very difficult, especially if one or both of the spiroplasmas react only partially with the test sera. INTERPRETATION OF RESULTS
Interpretation depends on the degree of knowledge about the mollicutes associated with the given host. Identification to the group level is usually not prob-
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lematic, if the unknown culture has a significant reaction to existing group antisera. Further testing, once preUminary screening has been carried out, consists of setting up complete dilution series with each of the components of the polyvalent sera. The final titer is determined by these tests. In many cases with tabanid spiroplasmas, some intergroup cross-reactivity, especially one-way crosses, has been observed. This circumstance, while unfortunate in one sense, can be exploited in defining spiroplasma serovars. For example, we always record titers of group VIII strains against all three representative group VIII sera. The set of three titers defines a level of reactivity that defines common serovars that occur in Georgia (French et al, 1990; Whitcomb et al., 1992a). Similarly, strains that react to polyvalent sera numbers 3 or 4 are tested against all four of the component sera of these polyvalent reagents. The matrix of titers obtained, as with group VIII, establishes four titers for each isolate and is a sensitive means of establishing serovar identity.
Discussion Mollicutes abound in arthropods, especially in insects. The meaning of infraspecific mollicute clusters from these vast reservoirs is not yet clear, but has formed a rich basis for speculation about the role that habitat, natural selection, and genetic drift play in the determination of mollicute serovars (Whitcomb, 1994). Identification of a large bank of strains related at various hierarchical levels will contribute to an understanding of the natural ecology of mollicutes in their insect hosts.
References French, F. E., Whitcomb, R. F., Tully, J. G., Hackett, K. J., Clark, E. A., Henegar, R. B., and Rose, D. L. (1990). Tabanid spiroplasmas of the southeastern USA: New groups, and correlation with host life history strategy. Zentralbl. BakterioL, Suppl. 20, 441-444. Gasparich, G., Whitcomb, R. F., French, F. E., Clark, E. A., and Tully, J. G. (1993). Serologic and genomic relatedness of group VIII and group XVII spiroplasmas and subdivision of spiroplasma group VIII into subgroups. Int. J. Syst. Bacteriol. 43, 338-341. Hackett, K. J., and Clark, T. B. (1989). Ecology of spiroplasmas. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.). Vol. 5, pp. 113-200. Academic Press, San Diego, CA. Le Goff, F., Humphery-Smith, I., Leclercq, M., and Chastel, C. (1991). Spiroplasmas from European Tabanidae. Med. Vet. Entomol. 5, 143-144. Markham, P., Clark, T. B., and Whitcomb, R. F. (1983). Culture techniques for spiroplasmas from arthropods. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 217-223. Academic Press, New York. Rodwell, A., and Whitcomb, R. F. (1983). Methods for direct and indirect measurement of myco-
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plasma growth. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 185-196. Academic Press, New York. Tully, J. G., Rose, D. L., Clark, E., Carle, P., Bove, J. M., Whitcomb, R. F., Colflesh, D. E., Henegar, R. B., and Williamson, D. L. (1987). Revised group classification of the genus Spiroplasma (class Mollicutes) with proposed new groups XII to XXII. Int. J. Syst. Bacteriol. 31, 357-364. Tully, J. G., Bove, J. M., Laigret, P., and Whitcomb, R. F. (1993). Revised taxonomy of the class Mollicutes: Proposed elevation of a monophyletic cluster of arthropod-associated mollicutes to ordinal rank (Entomoplasmatales ord. nov.), with provision for familial rank to separate species with nonhelical morphology (Entomoplasmataceae fam. nov.) from helical species {Spiroplasmataceae), and emended description of the order Mycoplasmatales, family Mycoplasmataceae. Int. J. Syst. Bacteriol. 43, 378-385. Tully, J. G., Whitcomb, R. F., Hackett, K. J., Rose, D. L., Henegar, R. B., Bove, J. M., Carle, P., Williamson, D. L., and Clark, T. B. (1994). Taxonomic descriptions of eight new non-sterolrequiring mollicutes assigned to the genus Mesoplasma. Int. J. Syst. Bacteriol. 44, 685-693. Whitcomb, R. F. (1983). Culture media for spiroplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.) Vol. 1, pp. 147-158. Academic Press, New York. Whitcomb, R. F. (1994). The species concept in eukaryotes and prokaryotes: Search for a synthesis. lOM Lett. 3, 1-7. Whitcomb, R. F., Bove, J. M., Chen, T. A., Tully, J. G., and Williamson, D. L. (1987). Proposed criteria for an interim serogroup classification for members of the genus Spiroplasma (Class Mollicutes). Int. J. Syst. Bacteriol. 37, 82-84. Whitcomb, R. F., French, F. E., Tully, J. G., Gasparich, G. E., Bove, J. M., Carle, P., Clark, E. A., and Henegar, R. (1992a). Tabanid spiroplasma serovars. lOM Lett. 2, 115. Whitcomb, R. F., Tully, J. G., Williamson, D. L., Bove, J. M., French, F. E., Konai, M., Gasparich, G., Abalain-Colloc, M. L., Saillard, C , Chastel, C , Carle, P., Rose, D. L., Henegar, R., Clark, E. A., and Hackett, K. J. (1992b). Revised classification of spiroplasmas. lOMLett. 2, 134. Williamson, D. L., and Whitcomb, R. F. (1983). Special serological tests for spiroplasma identification. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 249-259. Academic Press, New York.
SECTION
E
Experimental Infections
El INTRODUCTORY REMARKS Joseph G. Tully
Experimental infections with mollicutes have played both a historic and an important role in establishing the ability of these organisms to produce disease in a variety of hosts. Indeed, the first organism ever described for the group, the agent of contagious bovine pleuropneumonia (now termed Mycoplasma mycoides subsp. mycoides), was cultivated from an experimentally induced localized infection in a rabbit (Nocard et al., 1896). In the intervening years, other experimental infections with these organisms have provided crucial evidence of their pathogenicity, as well as noteworthy knowledge regarding a variety of host responses. The early advances were accomplished despite the then unknown obstacles of mixed microbial agents in the inocula (including other mollicutes) and the presence of other latent infections (again, those due to mollicutes) in the hosts employed. The chapters that follow in this section demonstrate some of the meaningful advances that have taken place both in the application and in the information gained from current experimental infections with mollicutes. Experimental infections can define the mechanism of attachment of mollicutes to target cells and the components on the organism that promote infection. Likewise, a variety of clinical, microbiologic, and immunological responses to infection can be determined, and the efficacy of vaccines and antibiotic agents established. However, several important common themes are apparent in these chapters. The results of any experimental model are very dependent on the status of the host or on the cell line in those in vitro techniques. Obviously, the host employed should be free of latent mollicutes, or at least the presence of concurrent mollicute flora should be clearly defined. Intensive surveillance with appropriate detection methods can usually determine colonization with, or exposure to, 325 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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mollicutes. Pathogen-free or cesarian-derived vertebrates are usually preferred, but not always available for some hosts. The presence of other microbial agents (viruses, etc.) in the host can also profoundly affect the outcome of the experimental challenge. Strain variation within host species can also have a dramatic effect on the response and results of an experimental infection, as can age of the host and a variety of environmental conditions (temperature, etc.). Strain variation within the challenge organism has been well documented to have a major impact on the ability to induce disease, and only a few passages of the organism outside the natural host can rapidly attenuate the challenge culture. Different routes of challenge and the number of organisms in the challenge will also condition host responses. As with laboratory studies on mollicutes, any decision to perform experimental infections, regardless of the host involved, should be made with a consideration of the potential hazards of such challenges and the institution of proper containment. Dissemination of infectious aerosols has occurred both within animal colonies and in laboratory personnel, the latter involving an experimental animal challenge with a human respiratory pathogen. Cell culture infections, particularly with mollicutes and viruses transmissible to humans (such as HIV) or animals, should be performed in appropriate biohazard facilities. All experimental challenges in animals should be carried out with respect to the humane treatment and welfare of such hosts. Many of the experimental infections detailed in the earlier volume of this series (Tully and Razin, 1983) are still relevant, especially in regard to those specialized techniques that have not materially changed. The current approach has been to update experimental infections where new information might have become available on a challenge organism, a particular host, or in the methodology to assay host response. Finally, it should be noted that animal models do not always replicate natural disease, but that much new knowledge can be gained from such experimental studies.
References Nocard, E., Roux, E. R., with Borrel, Salimbeni, and Dujardin-Beaumetz, (1896). Le microbe de la peripneumonie. Ann. Inst. Pasteur, Paris 12, 240-262; for English translation see Rev. Infect. Dis. 12, 354-358 (1990). Tully, J. G., and Razin, S., eds. (1983). "Methods in Mycoplasmology," Vol. 2, pp. 440. Academic Press, New York.
E2 EXPERIMENTAL MYCOPLASMAL RESPIRATORY INFECTIONS IN RODENTS Gail H. Cassell and A. Yancey
Introduction Experimental production of respiratory disease in animal models is essential for the fulfillment of Koch's postulates in establishing the etiologic significance of a particular mycoplasma species. In addition, relevant animal models can play a pivotal role in elucidating host and microbial factors involved in disease pathogenesis. Use of laboratory rodents offers many advantages to larger animals, including nonhuman primates. With respect to mycoplasmas, the availability of a large number of inbred strains of rats and mice with defined microbiological and environmental backgrounds is the most important advantage since heredity, synergism with other infectious agents, and exposure to environmental ammonia and oxidants are known to influence greatly disease character and severity (Cassell et al., 1985). The availability of transgenic rats and mice and immunocompromised rats and mice offers great opportunities for further defining the role of immunologic factors both in protection against the disease and in disease pathogenesis. Small laboratory animals have been utilized to establish highly reproducible models of several mycoplasmal respiratory diseases (Table I). While guinea pigs, hamsters, and cotton rats have been useful in initially establishing the pathogenic potential of Mycoplasma pneumoniae (Jacobs et al., 1988; Jemski et al., 1977), further work with these animal species is somewhat limited by the lack of availability of immunologic reagents. Newborn mice have been used to document the invasive potential of M. pneumoniae and occurrence in extrapulmonary sites, but mice and rats in general have been underutilized as models of M. pneumoniae infection. Newborn mice have also been valuable in establish327 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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Gail H. Cassell and A. Yancey TABLE I SMALL LABORATORY ANIMALS ESTABLISHED AS MODELS OF MYCOPLASMAL RESPIRATORY DISEASES
Animal
Mycoplasma species
Disease(s)
Guinea pig Hamster Cotton rats Rats Mice
Mycoplasma pneumoniae M. pneumoniae M. pneumoniae M. pulmonis M. pulmonis
Mice Mice
M. pneumoniae Ureaplasma urealyticum
Pneumonia Pneumonia Pneumonia Rhinitis, tracheitis Rhinitis, tracheitis, otitis, pneumonia Pneumonia Pneumonia
Natural host(s) Humans (all ages) Humans (all ages) Humans (all ages) Rats and mice Rats and mice Humans (all ages) Humans (newborn infants)
ing the pathogenic potential of Ureaplasma urealyticum in the production of pneumonia in newborn human infants (Rudd et al., 1989). Germ-free rats and mice have been invaluable in establishing the etiologic significance of M. pulmonis in naturally occurring chronic respiratory diseases of laboratory rats and mice (Cassell et aL, 1986).
Materials For Inoculation Freshly thawed mycoplasma suspension diluted in complete mycoplasma medium Sterile mycoplasma medium for inoculation of control animals Suitable pathogen-free animals maintained under germ-free conditions or maintained in sterile Isocages with sterile hardwood chip bedding and provided ad libitum with sterile food and water and maintained with intracage ammonia <25 ppm (0.025 ml/liter) Tuberculin syringes, sterile, 1 ml with 25-gauge needles for the administration of anesthesia Pentobarbital sodium (0.06 mg/g) for anesthesia or alternative anesthetic approved for use in small laboratory animals Eppendorf 25-|xl pipetter with sterile, disposable tips for intranasal inoculation Lamp with flexible neck (60- or 100-W bulb)
For Collection of Samples Sterile surgical instrument pack: 2-inch x 2-inch gauze sponges; 4-inch x 4-inch gauze sponges; mosquito hemostats; straight Mayo scissors; suture wire scis-
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sors; suture thread, cut into 3- to 4-inch lengths; Iris scissors; Adson forceps; and dissecting pins Dissecting board, covered with absorbent pad Glass beakers, 500 ml (one containing H2O and one containing absolute ethanol for washing and flame disinfection of instruments between samples) Absolute alcohol for decontamination of fur and skin; 95% cold ethanol or 10% PBS-buffered formalin for fixation of tissues Burner for disinfection of instruments Tissue grinders: mortar and pestles or tissue homogenizers, either mechanically or electrically driven plunger and shaft Nasopharyngeal cotton swabs Syringes with 18- to 25-gauge needles, sterile, 1 cc (depending on animal size) Sample collection fluid: sterile phosphate-buffered saline (PBS) with 2% fetal calf serum (FCS), pH 7.4; or mycoplasma broth Tubes for blood collection at time of exsanguination Anesthetic: sodium pentobarbital or alternative Surgical clippers for hair removal
Procedure Inoculation
1. Anesthetize the animal with 0.06 mg/g body weight of sodium pentobarbital injected intraperitoneally (ip). 2. When anesthetized, slowly deliver 25 U | L1 of organisms per nostril with the animal held with its head in an upright position. 3. When all the inoculum has been inhaled, place animals in a lateral position and keep warm with lamps until the animals have recovered (approximately 1 hour).
Sampling PRINCIPLES OF SPECIMEN COLLECTION Euthanasia and Bleeding
Animals can be humanely killed by asphyxiation with CO2, by cervical dislocation, or preferably by an overdose of pentobarbital sodium. Exsanguination decreases the chances of contamination of culture specimens by blood. When
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deep general anesthesia is attained, place the animal on its back and incise either the axial or inguinal area until the artery is severed. Blood accumulates in the pocket thus formed and can be transferred to a suitable tube. This method avoids the vagaries of cardiac bleeding as well as possible contamination of the thoracic cavity. Sampling Sites
Until the optimum body site for mycoplasmal detection [either by culture or polymerase chain reaction (PCR)] has been established, the following sites within the respiratory tract should be sampled: nasal passages, nasopharyngeal tube, oropharynx, larynx, middle ears, and lungs. Optimum detection is achieved by routine sampling of multiple sites. Methods of Specimen Collection for Detection of Mycoplasmas in the Respiratory Tract
Proper collection of appropriate specimens is as important as preparation and quality control of media or other procedures used for detection. Mycoplasmas most commonly reside on the ciliated epithelial cells within the respiratory tract and are often unevenly distributed. Therefore, lavage is one of the better sampling techniques. Swabbing of the mucosal surfaces also is effective but serves best for more regional sites or single organs (e.g., larynx). Tissue homogenates are usually less desirable except for special purposes, such as quantitative cultures, but may be used if appropriate measures are taken to prevent the effects of tissue inhibitors. The following stepwise procedure is designed for the comprehensive sampling of the respiratory tract; less complete protocols may be developed, depending on specific objectives. Instruments must be sterilized between each step by washing in water, dipping in ethanol, and flaming. If PCR is the detection method being used, even more extreme caution must be used and new instruments will be required for each site. 1. Clip the hair on the ventral surface of the animal from the chin to the pubis. 2. Place the animal in the supine position on a dissection board and secure the feet with pins. 3. Wet the entire ventral surface of the animal with absolute alcohol. 4. Grasp the skin over the larynx with forceps and excise with scissors. 5. The submaxillary salivary glands and lymph nodes will then be seen embedded in fat in the middle of the exposed field. 6. Lift the salivary glands and lymph nodes, excise them and the associated fat.
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7. The thin omohyoid muscle is then seen extending from the base of the tongue to the sternum, closely investing the larynx and trachea. 8. Lift the omohyoid muscle and excise it, exposing the larynx and trachea. 9. Grasp the larynx and insert a needle through the tracheal wall toward the major bronchi to obtain a tracheobronchial lavage specimen. For an adult rat, hamster, or guinea pig use a 20-gauge needle with a tuberculin syringe containing 0.6 ml collection fluid. For a mouse, use a 22-gauge needle and a tuberculin syringe containing 0.4 ml collection fluid. Express the collection fluid and, with the bevel of the needle down against the wall of the trachea, aspirate while moving the needle backward toward the larynx. Repeat two to five times. Greater volumes of fluid can be recovered by elevating the posterior part of the animal's body. 10. Remove the lower jaws carefully, dissecting all soft tissue attachments from the ventral surface of the larynx while avoiding contamination of the pharynx or lumen of the larynx. 11. Lift the larynx with forceps and swab the interior walls. Make sure the swab enters the larynx and not the esophagus. 12. A separate swab is inserted anteriorly into the nasopharyngeal tube. (This is not practical for mice as the nasopharyngeal tube is too small. Lavage with a small gauge needle is satisfactory.) 13. The soft tissue of the nose should be cut away with sterile scissors in order to reduce the number of bacterial contaminants. A needle is inserted into the nasopharyngeal tube and the nasal passages are flushed anteriorly into a collection tube. Recovery of collection fluid is enhanced by lowering the animal's head relative to the remainder of the body. 14. The tympanic bullae are exposed and swabbed with ethanol. A needle mounted on a tuberculin syringe is used to drill into the middle ears. Collection fluid is then alternately expressed and aspirated a few times in order to obtain a lavage specimen. 15. Specimens should be immediately processed for the culture of mycoplasmas as well as potential indigenous pathogens which could confound the interpretation of results.
Histopathology and Lesion Scoring
1. Gently reinflate the lungs with either cold 95% ethanol or buffered formalin. 2. Remove the needle and tie off the suture thread so that the lungs remain distended. 3. Remove the lungs from the thoracic cage and immerse in fixative (95% alcohol or buffered formalin) for at least 24 hours. 4. After fixation, remove the trachea and section longitudinally.
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5. Separate each lobe of lung and cut so that a single section can be obtained with the main stem bronchi cut in a longitudinal section. 6. After 48 to 72 hours of fixation, use 10% formic acid, or alternative, for demineralization. When the bone has softened, take three sections from the nasal passages: from a transverse plane immediately behind the developing incisors; at the level of the third palatine ridge; and from a transverse plane bisecting the orbit. This will give sections through all levels of the nasal passages, middle ears, and Harderian glands.
Discussion In order to assess appropriately the disease-producing potential of a mycoplasma in an experimental animal, many intrinsic and extrinsic factors, in addition to the virulence of the test organism, must be taken into consideration in experimental design. Selection of Inoculum As it is the case with other infectious agents, a wide variation in the virulence of different strains of mycoplasmas may exist. In addition, it is now apparent that certain surface antigens undergo a high rate of variation within a strain, often resulting in a mixed population containing organisms with different pathogenic potential. It is imperative that any inoculum to be used to infect animals be of the lowest number of artificial passages as possible and at the same time be well characterized. Ideally, the inoculum should be derived from a filter-cloned stock. It may also be wise to rapidly passage the clinical isolate several times in the laboratory animal to be used for development of the model so as to adapt the organism to the host better. If the purpose of animal inoculation is to clarify etiologic significance, simultaneous comparison of several isolates from diseased tissue is advisable. The number of organisms required for the establishment of colonization and disease will vary, as with other infectious agents, depending on the virulence of the infecting organism, the genetic background of the host, and environmental factors. Because of the overall low virulence of most mycoplasmas, a relatively large number of organisms (10^) should be used initially. Ideally, for characterization of the model the biological parameters of the 50% infective dose, the 50% gross pneumonia dose and microscopic lesion dose, and the 50% lethal dose should be calculated by the method of Reed and Muench (1938). For further comparison of organism virulence, genetic susceptibility of the host, or other
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variables, the 50% dose should be used as well as one dose above and one below the 50% dose. Administration of Inoculum
One of the most striking features of experimentally induced mycoplasma respiratory disease is the extreme variability in the incidence, severity, and extent of both upper and lower respiratory tract lesions after experimental infection via intranasal inoculation. This variability is seen even when care is taken to minimize the effects of intrinsic and extrinsic factors such as age, genetic constitution, and levels of intracage ammonia, all of which are known to markedly influence disease incidence and severity. One of the major sources of variation in intranasal inoculation is the proportion of the inoculum which actually reaches the lungs. This proportion can be affected by such factors as the depth and type of anesthesia and the volume inoculated. Lightly anesthetized animals often swallow part of the inoculum; in addition, many anesthetics affect pulmonary defense mechanisms. In hamsters intranasally inoculated with 2, 20, or 200 |JL1 of M. pneumoniae (with the concentration adjusted to give the same total dose per animal), only the largest volume resulted in the deposition of the organisms in the lungs (Jemski et al., 1977). At the same time, administration of excess liquid must be avoided to prevent drowning. The use of a small-particle aerosol (with a mass median diameter of 2.3 |xm) results in more uniform deposition of mycoplasmas in both the upper and the lower respiratory tracts. However, considerable variability is still observed between animals in the numbers of lung lobes involved and the severity of disease present. In addition, the expense and time required for establishment and characterization of aerosol models make them impractical for routine use. Aerosol models are thus used to address selective questions, e.g., those related to pulmonary clearance. The methods used in establishment and characterization of aerosol models in laboratory rodents have been described in detail elsewhere for both M. pulmonis (Davis et al, 1986) and M. pneumoniae (Jemski et al., 1977). Requirement for Histopathological Examination
One should be aware that extensive microscopic changes can be present following exposure of the respiratory tract to mycoplasmas, even in the absence of overt clinical signs and symptoms and even in the absence of gross lesions at autopsy. Therefore, rigorous histopathologic examination of infected animals is required for assessment of disease-producing potential. Interpretation should be based on detailed comparison with the morphology of matched broth-inoculated control animals. The severity of lesions in the nasal passages and lungs can be scored either subjectively (Davis et al., 1985) or morphometrically (Davis et al.,
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1982). Lesion severity in the trachea and middle ears is scored subjectively. Data should be statistically analyzed using appropriate tests for nonparametric data, i.e., subjective lesion scores. Synergism with Other Infectious Agents and Environmental Impact Because respiratory disease research involving any infectious agent is easily compromised by environmental factors and indigenous infections (Bhatt et al., 1986), the primary requirement of an animal model should be the use of environmentally and microbiologically defined animals. Indigenous respiratory infections, many of which are clinically silent, can have subtle effects on nonspecific pulmonary clearance, immunological responses, and respiratory physiology. Mice and rats are two of the few groups of laboratory animals in which the microbial flora (both commensal and pathogenic) of the respiratory tract are well defined (Bhatt et al, 1986). The morphology of the lungs of normal rats and mice is well characterized, as are the histopathologic changes associated with naturally occurring murine pathogens. Most importantly, pathogen-free rats and mice are available and can be maintained to ensure their pathogen-free status (Bhatt ^r a/., 1986). It is important to note that a number of mycoplasma species can be commonly isolated from the respiratory tracts of normal laboratory rodents. Because mycoplasmas that normally colonize the genital tracts of rats and mice can be transmitted in utero, mere cesarian derivation and barrier maintenance does not guarantee the eradication of indigenous mycoplasmas or, for that matter, other rodent pathogens. Hence appropriate surveillance systems must be used over long periods to carefully document the pathogen-free status of a given colony (CasselUra/., 1986). When using laboratory rodents to establish the etiologic significance of mycoplasmas in respiratory diseases, the impact of environmental factors should be taken into account. For example, intracage ammonia levels (even as low as 25 ppm, levels common in most vivariums) can enhance the severity of respiratory disease produced by M. pulmonis (Cassell et al, 1986). Animal weight and numbers per cage, type of bedding, and frequency of cage changing can all modify intracage ammonia levels. We have also previously shown that exposure of newborn mice to 80% oxygen, a concentration used commonly in human neonatal respiratory care, results in persistence of U. urealyticum in the lungs of newborn mice, potentiates the inflammatory response, and turns an otherwise self-limiting pneumonia into a lethal disease (Crouse et al, 1990). Heredity, Age, and Gender Depending on the genetic background of the host, the inflammatory response and disease severity can vary greatly. For example, intranasal inoculation of
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C3H/HeN mice with the same strain of M. pulmonis in comparable numbers of organisms per gram of lung weight, as used in rats, results in an acute fatal disease in mice characterized by pulmonary edema and hemorrhage compared to a slowly progressing chronic bronchopneumonia when inoculated into highly susceptible Lew rats (Cassell et aL, 1985, 1986). In contrast, the same strain of M. pulmonis inoculated intranasally into C57B1/6N mice reared and maintained under identical microbiological and environmental conditions causes a mild, self-limiting acute bronchopneumonia to develop. When attempting to establish a reproducible animal model, inclusion of sufficient animals of different ages and sex is also important. For example, U. urealyticum isolated from pneumonic lungs of human infants can produce a similar disease when inoculated intranasally into newborn mice but not 14-dayold mice (Rudd et al., 1989). Data also indicate that when all other variables are comparable, male mice are more susceptible to M. pulmonis-inducQd respiratory disease than female mice.
Disease Progression Determining the exact number and timing of observation points following exposure of the respiratory tract to mycoplasmas is also an important consideration in establishing an optimal animal model. Depending on the species of mycoplasmas and the genetic background of the host, lesions may peak as early as 3-5 days or as late as 60 days postinoculation. Naturally occurring mycoplasmal respiratory diseases in animals are characterized by colonization of the entire respiratory tract followed by development of slowly progressing, chronic inflammation (Cassell et aL, 1985). For example, in F344 rats, microscopic lesions do not become evident for 14 to 28 days following intranasal inoculation of M. pulmonis, with peak lesions developing at 30 to 60 days. When initially establishing an animal model it is therefore important to include multiple observation points.
References Bhatt, P. N., Jacoby, R. O., Morse, H. C , and New, A. E., eds. (1986). "Viral and Mycoplasmal Infections of Laboratory Rodents: Effects on Biomedical Research." Academic Press, Orlando, FL. Cassell, G. H., Clyde, W. A., and Davis, J. K. (1985). Mycoplasmal respiratory infections. In "The Mycoplasmas" (S. Razin and M. F. Barile, eds.). Vol. 4, pp. 65-106. Academic Press, Orlando, FL. Cassell, G. H., Davis, J. K., Simecka, J. W., Lindsey, J. R., Cox, N. R., Ross, S. E., and Fallon, M. (1986). Mycoplasma infections: Disease pathogenesis, implications for biomedical research, and control. In "Viral and Mycoplasmal Infections of Laboratory Rodents: Effects on
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Biomedical Research" (P. N. Bhatt, R. O. Jacoby, H. C. Morse III, and A. E. New, eds.), pp. 87-130. Academic Press, Orlando, FL. Crouse, D. T., Cassell, G. H., Waites, K. B., Foster, J. M., and Cassady, G (1990). Hyperoxia potentiates Ureaplasma urealyticum pneumonia in newborn mice. Infect. Immun. 58, 34873493. Davis, J. K., Thorp, R. B., Maddox, P. A., Brown, M. B., and Cassell, G. H. (1982). Murine respiratory mycoplasmosis in F344 and LEW rats: Evolution of lesions and lung lymphoid cell populations. Infect. Immun. 36, 720-729. Davis, J. K., Parker, R. F., White, D., Dzicdzic, D., Taylor, G., Davidson, M. K., and Cassell, G. H. (1985). Strain differences in susceptibility to murine respiratory mycoplasmosis in C57BL/6 and C3H/HeN mice. Infect. Immun. 50, 647-654. Davis, J. K., Thorp, R. B., Parker, R. F., White, H., Dziedzic, D., D'Arcy, J. D., and Cassell, G. H. (1986). Development of an aerosol model of murine respiratory mycoplasmosis in mice. Infect. Immun. 54, 194-201. Jacobs, E., Stuhlert, A., and Drews, M. (1988). Host reaction to Mycoplasma pneumoniae infections in guinea pigs preimmunized systematically with the adhesin of this pathogen. Microb. Pathog. 5, 259-265. Jemski, J. V., Hetsko, C. M., Helms, C. M., Grizzard, M. B., Walker, J. S., and Chanock, R. M. (1977). Immunoprophylaxis of experimental Mycoplasma pneumoniae disease: Effect of aerosol particle size and site of deposition of M. pneumoniae on the pattern of respiratory infection, disease, and immunity in hamsters. Infect. Immun. 16, 93-98. Reed, L. J., and Muench, H. (1938). A simple method of estimating 50 percent endpoints. Am. J. Hyg. 27, 493-497. Rudd, P. T., Cassell, G. H., Waites, K. B., Davis, J. K., and Duffy, L. B. (1989). Ureaplasma urealyticum pneumonia: Experimental production and demonstration of age-related susceptibility. Infect. Immun. 57, 918-925.
E3 UROGENITAL INFECTIONS IN RODENTS Patricia M. Furr and David Taylor-Robinson
Introduction As a group, the Rodentia comprises a large number of well-known species as well as some lesser known ones, but for the purposes of this chapter, only animals used commonly as models for infection will be considered, namely mice, rats, guinea pigs, hamsters, and rabbits. Since naturally occurring mycoplasmas should be excluded prior to the use of these animals for experimental infection, consideration will be given to such organisms and the techniques for their isolation as well as to those introduced experimentally. The procedures required for the experimental inoculation of the genital tract of these small animals are outlined, although these methods are suited equally to examination of other anatomical sites and of larger animals. Mycoplasmas tend to be species specific and not to cross major animal species barriers naturally. However, the use of sex hormones is known to predispose mice and hamsters to genital tract colonization with mycoplasmas of human as well as murine origin. The procedures required to induce such susceptibility are described.
Materials and Procedures Media Culture results are a reflection of the quality of the medium used and it will probably never be possible to produce a medium in which it is possible to culture 337 Molecular and Diagnostic h e d u r e s in Mycoplasmology, Vol. I1
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every mycoplasmal species. SP4 medium (Tully et al., 1979; see also Chapter A2 in Vol.1) is more sensitive for the isolation of some mycoplasmas than other media but, routinely, we have used "Edward" broth medium (Freundt, 1983) for primary isolation and for culturing mycoplasmas for experimental inoculation of animals. A liter of this medium is composed of PPLO broth base (Difco) (2.1%, w/v), 700 ml; benzylpenicillin (Glaxo) (1000 lU/ml), 10 ml; thallous acetate (British Drug Houses) (2.5%, w/v), 20 ml (sometimes omitted or reduced in concentration, especially when culturing ureaplasmas); yeast extract (Distillers' Co.) (25% w/v), 100 ml; horse serum (Imperial Laboratories), 200 ml; and phenol red (BDH) (1%), 20 ml. This is regarded as a standard liquid medium (SLM) which is suitable for transportation and storage of specimens and to which may be added one of the following three substrates for culture of mycoplasmas; glucose (BDH) (10%, w/v), 10 ml; L-arginine hydrochloride (Sigma) (10%, w/v), 20 ml; or urea (BDH) (10%, w/v), 10 ml. The pH of each medium is adjusted by the addition of normal sodium hydroxide or normal hydrochloric acid to pH 7.8 for glucose-containing, 7.0 for arginine-containing, or 6.5-6.8 for urea-containing media, respectively. Swab Specimens
Plain cotton swabs are preferable to calcium alginate or serum-coated swabs because they are cheaper to purchase and, more important, contain fewer inhibitors (P. M. Furr and D. Taylor-Robinson, unpublished observations). For collection of vaginal swab specimens or, indeed, throat swab specimens from mice, a nasopharyngeal swab (MW 142; Medical Wire and Equipment Co.) is used. It is feasible to use a throat swab (MW 104) at either site in larger animals such as guinea pigs. The swab is inserted and rotated to abraid epithelial cells and the contents of the swab are expressed in the appropriate broth (1.8 ml, for example) by squeezing the swab against the side of the vial. The swab is then discarded as the stick or cotton wool may contain resins which are toxic for the mycoplasmas. This sample, now regarded as a 10"^ dilution, should be transported rapidly, preferably on wet ice, to the laboratory and cultured or stored at -70°C until required. Vaginal Washings
These specimens are collected for two different reasons which govern the choice of fluid with which the vagina is washed. The first is to undertake mycoplasmal culture, for which either SLM or a complete mycoplasmal medium is used. The second is to assess the stage of the murine reproductive cycle, when isotonic saline or phosphate-buffered saline is used (this latter procedure is
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described in detail later). Vaginal washing is achieved by introducing 50 JJLI of fluid into the vagina using a Finnpipette (Labsystems, Jencons), withdrawing the fluid, and reintroducing it twice more before collection. The volume of fluid used is determined by the size of the animal and may be increased to 100 |JL1 for a fully adult mouse. Repeat samples may be obtained at regular intervals from the same mouse and there is evidence, from introducing methylene blue dye (P. M. Furr, personal observation), that the fluid does not pass the cervix. Upper Genital Tract Washings
Washings may be taken from the uterus at postmortem examination using a technique similar to that described for the perfusion of mouse lungs (TaylorRobinson et aL, 1972). By introducing a small cannula through the cervix and clamping this in place, the uterine horns may be aspirated with broth for the isolation of mycoplasmas, or histological fixative may be introduced. Tissue Samples
In order to examine genital tissues by cultural or histological techniques, it is necessary sometimes to sacrifice animals. The animal is killed by administration of a lethal dose of barbiturate or by cervical dislocation in the small animal species. Exsanguination is undertaken preferably by cardiac puncture, although, in the mouse, cutting blood vessels in the axilla may be preferred. The fur is swabbed with 70% ethanol and allowed to dry and an incision is made into the body cavity using sterile instruments, with aseptic technique being observed throughout. The tissue samples, such as uterine horns and ovaries, are removed and divided: one piece being placed in a preweighed sterile container and the other in an aliquot of formal saline or other fixative if histology and/or electron microscopy are required. Samples for culture are weighed and a 10% (w/v) suspension, regarded as a 10"^ dilution, is prepared by mascerating the tissue in a Ten Broeck tissue grinder. If electrically operated blenders are used, care should be taken to keep the vessel containing the tissue in wet ice, as the heat generated may have a deleterious effect on the mycoplasmas. Isolation of Mycoplasmas and Estimation of Numbers
In order to determine the existence and number of mycoplasmal organisms present in a specimen, a series of 10-fold dilutions is prepared from the (10~ 0 dilution. This may be done in any volume, but 0.2 ml diluted in 1.8 ml of the required medium up to a dilution of 10"^ is convenient. The dilutions are contained in screw-cap vials of 2.5 ml capacity. Air-tight caps are required to
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prevent diffusion of gases from the medium causing nonspecific color changes. The vials are incubated aerobically at 37°C and observed for an alteration in the pH of the medium as denoted by a color change from red to yellow in glucosecontaining medium or from yellow to red in arginine- or urea-containing media. This denotes mycoplasmal growth and the last vial in which it occurs is deemed to contain 1 color-changing unit. If quantitation is not required and it is necessary only to see if mycoplasmas are present, it is still advisable to dilute the sample to at least 10~^. The reasons for this are several, as outlined elsewhere (TaylorRobinson, 1989), with diluting inhibitors being a principal one. Following experimental inoculation of animals with a known mycoplasma, it may not seem reasonable to confirm that all color changes are due to mycoplasmal growth. The situation is different, however, in a screening program where the existence of a mycoplasma is usually confirmed by seeking colonies on agar medium. For this purpose, mycoplasmal agar (Oxoid) (1%, w/v) is prepared in the PPLO broth base and the remaining components, as described earlier, are added aseptically. Agar plates are incubated under microaerophilic conditions (5% CO2 + 95% N2) normally at 36.5-37.5°C. This is satisfactory for samples collected from mice, rats, and hamsters, but the body temperature of guinea pigs and rabbits is higher and cultures may benefit from being incubated at 38°-39°C. All cultures should be inspected regularly and not discarded as negative for at least 3 weeks. Some acholeplasmas, such as A. multilocale from guinea pigs, do not metabolize glucose or arginine and their presence should be confirmed by subculture from broth to agar medium at 2- to 3-day intervals. Strict anaerobic conditions are required for the isolation of the arginine-metabolizing murine species M. muris (McGarrity et al, 1983), and similar conditions are needed for a glucose-fermenting mycoplasma, resembling M. alvi, isolated from the gastrointestinal tract of wild mice (Gourlay and Wyld, 1976).
Hormone Treatment Hormone treatment has a key role in predisposing female mice to colonization of the genital tract by various mycoplasmas. Progesterone or estrogen has been used and those mycoplasmas that have so far been found to colonize under the influence of one or other of these hormones are shown in Table I. The hormones are given in the following regimes: 150 mg/ml progesterone (Depo-Provera) (Upjohn) is diluted 1:12 in isotonic saline and 0.2 ml (2.5 mg) is administered subcutaneously on four occasions at weekly intervals using a 1-ml syringe and 25-gauge x Vs inch needle, or 5 mg/ml estradiol benzoate (Intervet UK Ltd) is injected in a similar regime to that for progesterone, with each mouse receiving 0.1 ml (0.5 mg) subcutaneously on four occasions, at weekly intervals.
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TABLE I EFFECT OF HORMONES ON COLONIZATION OF GENITAL TRACT OF FEMALE B A L B / C MICE BY VARIOUS MYCOPLASMAL SPECIES
Colonization when treated with Mycoplasmal species
Natural host
Substrate metabolized
Terminal Structure
Progesterone
Estradiol
Mycoplasma hominis M. salivarium Ureaplasma urealyticum M. fermentans M. pirum M. penetrans M. pneumoniae M. genitalium'^ M. pulmonis M. neurolyticum M. gallisepticum Acholeplasma laidlawii
Humans Humans Humans Humans ?Humans Humans Humans Humans Murine Murine Avian Ubiquitous
Arginine Arginine Urea Glu/Arg Glu/Arg Glucose Glucose Glucose Glucose Glucose Glucose Glucose
+ + + + + + -
+ + + -
+ + + + + -
^ Also colonizes the genital tract of female hamsters when they are treated with progesterone.
Vaginal Cytology A nasopharyngeal swab is used to abraid the vaginal epithelial cells, as described previously. The swab is rolled along a 3 x 1-inch glass microscope slide to transfer the material for cytological examination, and the slide is left to dry at room temperature. The cellular material is fixed by immersing the slide in methanol for 30 minutes. After fixation, slides are transferred to freshly prepared Giemsa stain for a further 30 minutes and then they are washed briefly in 30% methanol, followed by buffered distilled water, pH 6.8, and finally deionized water. The smears are dried at room temperature and are observed microscopically using a 10 or 40x objective. The cellular contents of the slides are recorded and the stage of the murine reproductive cycle is determined by following the criteria of Rugh (1968), which are as follows: 1. Diestrus: almost exclusively polymorphonuclear leukocytes (PMNL) (Fig. 1). 2. Proestrus: PMNL and nucleated epithelial cells in approximately equal proportions. 3. Early estrus: clearly defined epithelial cells, some nucleated. 4. Estrus: almost exclusively enucleated squamous epithelial cells (Fig. 2).
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Fig. 1 . Smear from a vaginal swab showing many polymorphonuclear leukocytes characteristic of the diestrous stage of the murine reproductive cycle. Magnification: 76x.
5. Postestrus: large epithelial cells with translucent nuclei and PMNL in approximately equal proportions.
Screening for Indigenous Mycoplasmas Prior to Experimental Infection Screening to ensure that animals are free of mycoplasmal organisms, especially the species to be used for experimental infection, is obviously important. In doing so, it is essential to examine not only the genital tract but also other anatomical sites. An existing infection at an extragenital site might result in the spread of organisms to the genital tract or could render the mice refractory to genital colonization with the same mycoplasma (Furr and Taylor-Robinson, 1992). The animal species being used will largely determine the mycoplasmal species for which there should be screening since, as mentioned previously, most mycoplasmas are species specific. It should be remembered, however, that occasionally they cross the animal species barrier and are recovered from closely
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i
Fig. 2. Smear from a vaginal swab showing clumps of enucleated squamous epithelial cells characteristic of the estrous stage of the murine reproductive cycle. Magnification: 76x.
related animals. The various mycoplasmas isolated from rodents and the range of animals from which each has been recovered are shown in Table II as an indication of what might be expected when screening. The methods used in screening have been outlined earlier.
Experimental Genital Tract Infection Animals (mice are used as an example here) are used that are considered to be free of the mycoplasma to be inoculated as a result of the screening process. The stage of the reproductive cycle is checked as described previously and the introduction of organisms coincides with the second injection of hormone. The organisms, stored at -70°C, are thawed rapidly and kept on wet ice during the inoculation procedure. Each mouse is held on its back in the palm of one hand of the operator, and 50 |JL1 of the inoculum is introduced into the vagina using a Finn pipette. The mouse is held in this position for a further few seconds to reduce seepage of the inoculum from the vagina, and then returned to its box. The
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Patricia M. Furr and David Taylor-Robinson TABLE II
NATURALLY OCCURRING MYCOPLASMAL SPECIES ISOLATED FROM LABORATORY RODENTS
Indicated mycoplasma recovered from Mycoplasma
Mouse
Rat
Guinea pig
Mycoplasma pulmonis M. neurolyticum M. collis M. muris M. alvi M. arthritidis M. caviae Acholeplasma laidlawii A. covigenitalium "Bovine group 7" M. oxoniensis A. multilocale
+ + + + + +
+ + +
+
+ + +
Hamster
Rabbit
+ + + +
ability of the mycoplasmal species to colonize is assessed at intervals, perhaps weekly; thereafter, vaginal swabs are collected for isolation of organisms and for vaginal cytology. Mice are sacrificed at intervals throughout the experiment and/or at the termination to determine whether the mycoplasmas have disseminated to the upper tract. In addition, urine may be aspirated directly from the bladder to determine if urinary tract involvement has occurred.
Discussion The need to screen animals for indigenous mycoplasmas within and without the genital tract before experimental genital tract inoculation has been pointed out. Attempting to detect mycoplasmas by a cultural method in at least a proportion of animals to be used for experimental purposes would seem more logical than using a serological technique, such as an enzyme immunoassay (Cassell et ai, 1981), the greater specificity of which might lead to a failure to detect. Of the mycoplasmas that are likely to be encountered, M. pulmonis is most likely to break species barriers and is, in this sense, ubiquitous. It is regarded normally as a respiratory pathogen of mice and rats, but some strains are also arthritogenic and others have been recovered from the genital tract (Casillo and Blackmore, 1972). Deeb and Kenny (1967) isolated two strains from the oropharynx of New Zealand white rabbits, and Cassell and Hill (1979) reported the isolation of
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M. pulmonis from the nasopharynx, conjunctivae, and genital tract of guinea pigs and Chinese and Syrian hamsters. Of course, where animals of different species are housed in close proximity to mice harboring a respiratory pathogen, spread might be expected and the occurrence of the organism in other species might represent no more than transient contamination rather than persistent colonization or infection. Nevertheless, the need to exclude this possibility is obvious. M. arthritidis is another murine mycoplasmal species that is important to exclude when contemplating experimental infection of the genital tract, particularly as it may be present as a latent infection and manifest itself only when the animals are subjected to stress (TuUy, 1969). Since hormone treatment increases the susceptibility of mice to experimental infection of the genital tract, it is possible that such treatment, although not tested, could activate a latent mycoplasmal infection. M. pulmonis will infect the genital tract of a proportion of mice without prior hormone treatment, but treatment with progesterone greatly enhances susceptibility (Furr and Taylor-Robinson, 1984, 1993a). The effect of the progesterone is to arrest the reproductive cycle in diestrus, dominated by PMNL, while estradiol reverts it to the estrous stage characterized by enucleated squamous cells. In the case of other mycoplasmas, a sharp distinction has been drawn between those for which progesterone is required to initiate and maintain genital tract colonization in mice and those for which estradiol is required. The contrasting effects of the two hormones in mice have been summarized previously (Furr and Taylor-Robinson, 1993b) (Table I). As in mice, treatment with progesterone is required for genital tract colonization of female hamsters with M. genitalium (Furr and Taylor-Robinson, 1990). The specificity of hormone treatment for different mycoplasmal species has been highlighted by the fact that progesterone not only fails to enhance colonization of mice with M. hominis but eliminates preexisting infection with this mycoplasma. Likewise, estradiol treatment eliminates infection with M. pulmonis (Taylor-Robinson and Furr, 1990). Whether mycoplasmas that have not been found to colonize the genital tract require the reproductive cycle to be brought to a stage other than diestrus or estrus is a moot point. Apart from hormone treatment, there is evidence that at least two other factors may be important in successful colonization. The first is the strain of the animal species. In this regard, a difference in susceptibility was observed in the responses of different hormone-treated mouse strains to colonization with Ureaplasma urealyticum; the BALB/c strain was more susceptible than the TO or CBA strains (Furr and Taylor-Robinson, 1989a), implying that predisposition to colonization is, at least partially, under genetic control. The susceptibility to other mollicutes may be influenced in a similar way. Second, the number of subcultures a mycoplasmal strain has received in the laboratory prior to inocula-
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tion may affect susceptibility. This was observed with M. pneumoniae when an isolate that had been passed five times in medium was found to colonize the genital tract of a larger proportion of progesterone-treated mice and for a longer period than when the FH type strain was used, a strain known to have undergone numerous passes in the laboratory (Furr and Taylor-Robinson, 1993b). Similarly, the multiply passed PG21 strain of M. hominis failed to colonize estradioltreated mice, whereas strains receiving few passes did colonize (Furr and TaylorRobinson, 1989b). The ability to swab the vagina easily and on repeated occasions is the major reason for using female animals. Repeated and even single swabbing of the male urethra is technically difficult unless much larger animals than mice, for example, rabbits, are used. Collection of urine samples from some male mice at the time of micturition is feasible (Taylor-Robinson and Furr, 1986), but is not consistently reliable. It should be pointed out that although vaginal swabbing provides information about the occurrence of genital tract colonization, recovery of organisms in this manner does not necessarily mean that the vagina is colonized. Indeed, the results of immunoelectron microscopy provide no evidence for attachment of either M. pulmonis or M. hominis organisms to vaginal epithelial cells, whereas attachment to the cervix and sometimes to the epithelium of the higher genital tract could be seen (Furr et al., 1995). Inevitably, swabs introduced into the vagina will abraid the cervix and also absorb epithelial cells which have sloughed from the upper tract. Furthermore, as an inoculum introduced intravaginally apparently does not pass the cervix, the presence of the mycoplasmas in the upper genital tract must signify an ascending infection.
References Casillo, S., and Blackmore, D. K. (1972). Uterine infections caused by bacteria and mycoplasma in mice and rats. J. Comp. Pathol. 82, 477-482. Cassell, G. H., and Hill, A. (1979). Murine and other small animal mycoplasmas. In "The Mycoplasmas" (J. G. Tully and R. F. Whitcomb, eds.), Vol. 2, pp. 235-273. Academic Press, New York. Cassell, G. H., Lindsey, J. R., Davis, J. K., Davidson, M. K., Brown, M. B., and Mayo, J. G. (1981). Detection of natural Mycoplasma pulmonis infection in rats and mice by an enzymelinked immunosorbent assay (ELISA). Lab. Anim. Sci. 31, 676-682. Deeb, B. J., and Kenny, G. E. (1967). Characterization oiMycoplasma pulmonis variants isolated from rabbits: 1. Identification and properties of isolates. J. Bacteriol. 93, 1416-1424. Freundt, E. A. (1983). Culture media for classic mycoplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, pp. 127-135. Academic Press, New York. Furr, P. M., and Taylor Robinson, D. (1984). Enhancement of experimental Mycoplasma pulmonis infection of the mouse genital tract with progesterone. J. Hyg. 92, 139-144. Furr, P. M., and Taylor-Robinson, D. (1989a). The establishment and persistence of Ureaplasma urealyticum in oestradiol-treated female mice. J. Med. Microbiol. 29, 111-114.
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Furr, P. M., and Taylor-Robinson, D. (1989b). Oestradiol-induced infection of the genital tract of female mice by Mycoplasma hominis. J. Gen. Microbiol. 135, 2743-2749. Furr, P. M., and Taylor-Robinson, D. (1990). The effect of sex hormones on mycoplasma genital infections. In "Recent Advances in Mycoplasmology" (G. Stanek, G. H. Cassell, J. G. Tully, and R. R. Whitcomb, eds.), pp. 223-226. Fischer, Stuttgart. Furr, P. M., and Taylor-Robinson, D. (1992). Mycoplasma pulmonis of the murine oropharynx protects against subsequent vaginal colonization. Epidemiol. Infect. I l l , 307-313. Furr, P. M., and Taylor-Robinson, D. (1993a). The contrasting effects of progesterone and oestrogen on the susceptibility of mice to genital infection with Mycoplasma pulmonis. J. Med. Microbiol. 38, 160-165. Furr, P. M., and Taylor-Robinson, D.(1993b). Factors influencing the ability of different mycoplasmas to colonize the genital tract of hormone-treated female mice. Int. J. Exp. Pathol. 74, 97-101. Furr, P. M., Sarathchandra, P., Hetherington, C. M., and Taylor-Robinson, D. (1995). Site of localisation of Mycoplasma pulmonis and Mycoplasma hominis in the genital tract of female mice demonstrated by culture, scanning- and immuno-electron microscopy. Int. J. Exp. Pathol. 76, 131-143. Gourlay, R. N., and Wyld, S. G. (1976). Ilsley-type and other mycoplasmas from the alimentary tracts of cattle, pigs and rodents. Proc. Soc. Gen. Microbiol. 3, 142. McGarrity, G. J., Rose, D. L., Kwiatkowski, V., Dion, A. S., Phillips, D. M., and Tully, J. G. (1983). Mycoplasma muris a new species from laboratory mice. Int. J. Syst. Bacteriol. 33, 350-355. Rugh, R., ed. (1968). "The Mouse. Its Reproduction and Development," pp. 38-39. Burgess, Minneapolis, MN. Taylor-Robinson, D. (1989). Genital mycoplasma infections. Clin. Lab. Med. 9, 501-523. Taylor-Robinson, D., and Furr, P. M. (1986). Urinary-tract infection by Mycoplasma pulmonis in mice and its wider implications. J. Hyg. 96, 439-446. Taylor-Robinson, D., and Furr, P. M. (1990). Elimination of mycoplasmas from the murine genital tract by hormone treatment. Epidemiol. Infect. 105, 163-168. Taylor-Robinson, D., Denny, F. W., Thompson, G. W., Allison, A. C., and Mardh, P.-A. (1972). Isolation of mycoplasmas from lungs by a perfusion technique. Med. Microbiol. Immunol. 158, 9-15. Tully, J. G. (1969). Murine mycoplasmas. In "The Mycoplasmatales and the L-Phase of Bacteria" (L. Hayflick, ed.), pp. 571-605. Appleton-Century-Crofts, New York. Tully, J. G., Rose, D. L. Whitcomb, R. F., and Wenzel, R. P. (1979). Enhanced isolation of Mycoplasma pneumoniae from throat washings with a newly modified culture medium. J. Infect. Dis. 139, 478-482.
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E4 EXPERIMENTAL MODELS OF ARTHRITIS Leigh Rice Washburn
Introduction The mycoplasmas as a group are probably the most common causes of naturally occurring arthritis in animals. Mycoplasmal diseases are economically important, with arthritis appearing either secondarily to infection at another site, such as the respiratory tract or mammary gland, or as the primary manifestation; the latter is particularly common in young animals (Cole et ai, 1985). In addition to their relevance in the field of veterinary medicine, the mycoplasmal arthritides have long served as models for human rheumatoid arthritis (RA), although the question of actual involvement of mycoplasmas in RA is unresolved. Despite sporadic reports over the past several decades of isolations of several Mycoplasma species, including the rat pathogen Mycoplasma arthritidis, from rheumatoid arthritic joints, the issue remains controversial. However, no etiologic agent has been discovered for this disease, and the mycoplasmas are well suited to such a role. Their complex nutritional requirements and close association with host tissue often render their isolation and cultivation a difficult and laborious process. Improvements in detection techniques and medium formulations may yet allow the identification of a mycoplasmal pathogen in RA. Several of the chapters in Section A (Cultivation and Morphology) of Vol. I are relevant to these investigations. While a role for mycoplasmas in RA is still in question, there is no doubt that they can cause other kinds of arthritis in humans. Arthritis has been reported as an occasional complication of M. pneumoniae respiratory infection, but most mycoplasmal arthritides are found in immunocompromised, especially hypogammaglobulinemic, patients and tend to involve species from the urogenital 349 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
Copyright © 1996 by Academic Press, Inc. All rights of reproduction in any form reserved.
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tract, such as M. hominis and Ureaplasma urealyticum. The experimental mycoplasmal arthritides may also provide excellent models for these human infections. The mycoplasmal arthritides occurring in small laboratory animals such as rats, mice, and, to a lesser extent, rabbits are those most often used as models for human disease. Highly homogeneous and genetically well-characterized strains of rats and mice are available in which individual aspects of the immune response can be precisely manipulated. The effect of these responses on arthritic manifestations of mycoplasmal infection and the effect of mycoplasmal infection and mycoplasmal products on the immune system can thus be very precisely characterized. Both of these are of critical importance in selecting a model for those human arthritides in which immunologic mechanisms of joint damage play a prominent role as well as those in which immunodeficiencies enhance the susceptibility of patients to systemic invasion. The following discussion is divided into two sections. The first is a description of the more common model systems, including animal strains, mycoplasmal species and dosages, routes of inoculation, and methods of assessment of illness. The second is a discussion of the use to which some of these models may be put, using examples drawn from the literature.
Materials and Procedures Previously, we described methods for the induction and assessment of arthritis in rats, mice, and rabbits (Cole and Washburn, 1983). These are reviewed here briefly and are updated. There are advantages and disadvantages to each model. Mice and rats are useful for several reasons: (i) They are economical to purchase and maintain; (ii) highly genetically defined strains are available; (iii) two mycoplasmal species found naturally in these animals are arthritogenic, M. arthritidis and M. pulmonis, and rats at least are susceptible to naturally occurring mycoplasmal arthritis; and (iv) the diseases induced in mice by both M. arthritidis and M. pulmonis are chronic and have been reported to include remissions and exacerbations (Cole et al., 1985). On the other hand, neither model precisely mimics any of the human arthritides, and little evidence exists at this time for an immunologic component, with the possible exception of the M. arthritidis superantigen MAM. Rabbits are more expensive, but an immune complex-mediated, chronic inflammatory arthritis can be induced by intraarticular injection that persists long after the causative mycoplasma agent has disappeared (Cole et al., 1985; Washburn and Ward, 1988). The disease mimics some of the immunologic aspects of chronic rheumatoid arthritis but does not include remissions and exacerbations and remains confined to the injected joint.
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Rats
M. arthritidis induces an acute, self-limited, systemic infection in rats that includes septic polyarthritis as a primary manifestation. Rat strains vary in susceptibility, and M. arthritidis strains, in virulence for rats, so both must be chosen carefully. Susceptible inbred rat strains include LEW, MWF, BS, BH, and DA (Binder et ai, 1990). In addition, the outbred Sprague-Dawley strain is quite susceptible and has been used extensively in the past. We use LEW rats in our laboratory because they are gentle, relatively inexpensive, and highly susceptible. WimlQUi M .arthritidis strains include Jasmin (ATCC 14124), JR3, and rat-passaged strains ISRl, 158plO, 158plOp9, 14124pl0, and 14152pl3. Virulence of other strains can be enhanced by serial passage through rats as previously described (Cole and Washburn, 1983). Rats weighing 120-200 g are injected intravenously in the caudal vein with 1 X 10^-10^ colony-forming units (CPU) of M. arthritidis in a total volume of 0.2-1.0 ml (Cole and Washburn, 1983). We use cultures that have been concentrated to contain 1 x lOiO-lO^i CFU/ml and stored at -70°C in mycoplasmal culture medium containing sucrose in place of serum to minimize the induction of antibodies against medium components. Large quantities of these stock cultures are prepared and stored frozen to minimize variability. Cultures are thawed just before use and adjusted to a concentration of 3.3 x 10^ CFU/ml; each animal receives 0.3 ml. For immunologic investigations, particularly when addressing the question of autoimmunity or antigenic mimicry, mycoplasmas should be grown in a dialyzate medium containing agamma horse serum or fetal bovine serum. The medium formulation used in our laboratory is described in a previous review (Washburn and Ward, 1988). Because disease develops rapidly in susceptible animals, especially with higher mycoplasmal doses, rats should be monitored daily for signs of arthritis and systemic illness during the first week, preferably at the same time each day to minimize the effects of diurnal variation. After that, they may be scored every 2 or 3 days, as needed, for the remainder of the observation period. Rats are scored as follows: Peripheral joints are examined and assigned a score of 0-4, depending on the relative severity of swelling; digits are examined separately and assigned a score of 1 each if any swelling is observed and 0 if it is not. In addition, we assign a score of 1 for partial and 2 for complete hind limb paralysis on each side. With high doses of virulent strains, joint swelling can appear as early as 1 day after injection, but signs usually appear between 2 and 4 days. With lower doses and less virulent strains, onset may be delayed several days. Because the scoring system is necessarily subjective, it is useful to include a second indicator of illness, such as weight loss, to supplement the data. Weight loss sometimes begins before the first signs of arthritis appear. It is also an
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indicator of systemic illness, whereas joint swelling measures only the arthritic manifestations of infection. In the LEW rats used in our laboratory, arthritis generally peaks in severity between 1 and 2 weeks after injection and drops rapidly after that. For immunization and M. arthritidis strain comparison studies, we usually use a 3-week observation period, but arthritis is rarely completely resolved by then, and much longer periods may be needed if complete resolution is desired or when studying a possible chronic phase. Any mycoplasmas isolated from injected rats should be reidentified as M. arthritidis. Methods for this include growth inhibition, epi-immunofluorescence, and colony blotting. The latter is our method of choice. It is performed by laying a piece of prewetted nitrocellulose membrane on the agar surface for 5 minutes and then staining the resulting colony replicas with species-specific antiserum exactly as is done for Western immunoblots. A problem is that M. arthritidis may be part of the normal flora of most rats and has been found even in barriermaintained animals (Cox et al., 1988). Although reliable strain markers are not yet available for M. arthritidis, we have observed restriction fragment length polymorphisms in chromosomal DNA from different strains. Determining whether isolates have the same patterns as the strain injected, while not conclusive, will provide at least presumptive evidence that they are one and the same. Differences in protein profiles on sodium dodecyl sulfate (SDS)-polyacrylamide gels or antigenic profiles on Western immunoblots between the strain injected and the samples recovered are not necessarily indicative of recovery of a different strain but may simply reflect antigenic variation within individual strains of M. arthritidis. Antibody responses can be monitored by routine serologic procedures, including enzyme-linked immunosorbent assay (ELISA), complement fixation, and Western immunoblotting, but not metabolism inhibition or growth inhibition assays because M. arthritidis does not include these antibodies in rats (Cole et al., 1985). Infection with M. arthritidis is characterized by an early and vigorous antibody response. We detected IgM antibodies by ELISA and Western immunoblotting in LEW rats as early as 3 days and IgG antibodies by 5 days after injection with 1.4 x 10^ CPU M. arthritidis strain 158plO (M. McKenzie and L. R. Washburn, unpublished observations). This appears to be a characteristic of infection with this agent and does not necessarily indicate previous exposure. Although studies on rat arthritis most often use M. arthritidis, M. pulmonis is also arthritogenic for rats, although less is known about differences in arthritogenicity for rats among M. pulmonis strains or susceptibility among rat strains. Techniques similar to those described earlier may be used to study this disease, although the selection of appropriate strains and routes of inoculation will have to be determined by the individual investigator. In one model, M. pulmonis derived from a naturally infected rat was introduced by intracerebral inoculation of 2 x
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10^-1.6 X 10^ CFU in 20 |xl into neonatal or intravenous inoculation of 1 x 10'^ CFU in 1 ml into adult Sprague-Dawley rats (Kohn and Chinookoswong, 1989). Joint swelling appeared within 3 to 8 days and persisted through 4 weeks postinoculation, which is the longest observation period reported for these studies. Mice Experimental arthritis can also be induced in mice with both M. arthritidis and M. pulmonis. Arthritis is assessed by a scoring system similar to that described for rats; in addition, we previously recommended the use of females for longterm studies because they tend to be less aggressive and less likely to incur injuries that could be confused with mycoplasma-induced lesions (Cole and Washburn, 1983). As with rats, mouse strains differ in susceptibility to M. arthritidis. Some susceptible strains include the outbred Swiss Webster and the inbred DBA, CBA, AKR, and C3H, whereas BALB/c and C57BL are more resistant. Mice bearing the H2^ and //2^ MHC haplotypes are also highly sensitive to M. arthritidis-induced toxic death, whereas H2^ mice are not (Cole et al., 1983); this may also affect experimental results. Toxicity is not necessarily related to the arthritogenic effects of M. arthritidis but is associated with the M. arthritidis superantigen MAM (Cole et al, 1983). Less is known about the relative virulence of different M. arthritidis strains for mice than for rats, but it is likely that strains exhibiting low virulence for rats are also less virulent for mice. Strains known to be arthritogenic for mice include PNZ9 (Kaklamani et al., 1991a) and the mouse-passaged 158plOp9 (Cole et al., 1971). Arthritis is most often induced by an intravenous injection of large numbers of mycoplasmas, 1 x 10^ CFU or more (Cole and Washburn, 1983), although intraperitoneal injection with similar doses has also been used (Cole et al., 1983). Arthritis develops in susceptible mice within 2-5 days after injection. Although most studies tend to focus on the acute stages of the disease. Cole et al. (1971) reported that a chronic arthritis, persisting through at least 38 weeks and characterized by periods of remission and exacerbation, could be induced by strain 158plOp9 in Swiss Webster mice. M. pulmonis is also capable of inducing a chronic migratory polyarthritis in mice; methods of induction and assessment of disease are similar to those used for M. arthritidis. As with M. arthritidis, results depend on both mouse and mycoplasmal strain (Cole et al., 1985). Sensitive mouse strains include A, DBA/2, AKR, C3H, C57BL, and AS/NIH. CBA mice are also susceptible but less so than C3H (Taylor et al., 1974). Younger animals, 4-8 weeks of age, are more susceptible than older animals (Harwick et al., 1973). Most studies have been done with the M. pulmonis JB strain of Barden and Tully (1969), which was originally isolated from an arthritic mouse joint. While other strains may show
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less arthritogenic potential, arthritogenicity can be enhanced by animal passage. Reported effective inoculum sizes vary between 1 x 10^ and 1 x 10^ CFU and should be determined for each M. pulmonis and mouse strain used. Arthritis is more readily induced by intravenous than intraperitoneal injection, whereas intranasal induces primarily respiratory disease (Harwick et al, 1973). Susceptibility is altered if mice are naturally infected with M. pulmonis or M. neurolyticum. Since these are relatively common murine pathogens, particularly M. pulmonis, mice should be carefully screened and maintained under appropriate conditions to avoid exposure. Rabbits A chronic inflammatory monoarticular arthritis can be induced in rabbits using either a single dose of viable M. arthritidis strain 158plOp9 (2 x 10^ CFU in 0.3 ml broth medium) injected intraarticularly into immunologically naive animals or a single dose of nonviable mycoplasmal antigen (1 mg mycoplasmal protein) injected intraarticularly into hyperimmunized animals. Arthritis is assessed by determining percentage increase in joint diameter measured by caliper. Detailed methodologies for these models have been published (Cole and Washburn, 1983; Washburn and Ward, 1988). Arthritis may also be induced by similar doses of M. pulmonis strain JB, although the chronic stage is not as severe or long-lasting. One of the most interesting aspects of this model is that viable mycoplasmas disappear from joint tissues within days after injection, whereas severe, active inflammation persists for several months.
Discussion Mycoplasmal arthritis models have been in use for many decades, and an extensive body of descriptive literature concerning these diseases exists (Cole et al., 1985). However, actual mechanisms of disease production have proven more elusive. A few examples of the more recent uses to which these models have been put and which have attempted to provide answers to that question are summarized next. Examining Arthritogenic Potential of Other Mycoplasmal Species Cedillo et al. (1992) have used the rabbit model described earlier to study the arthritogenic potential of the human respiratory pathogen M. pneumoniae. This is of interest not only because arthritis is an occasional complication of M. pneumoniae infection, but also because a mycoplasmal etiology for rheumatoid
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arthritidis cannot be ruled out at this time; mycoplasmas ordinarily found at other sites in the human body remain prime candidates for such an agent. In this study, 3 x 1 0 ^ CFU of M. pneumoniae (ATCC 29342) were injected intraarticularly into 2.5-kg female rabbits, resulting in an acute arthritis that persisted for 2 weeks. This model may prove useful for exploring the arthritogenic potential of other human mycoplasmal species as well. Localization of Mycoplasmas in Joint Tissues of Experimental Hosts
Methods for the detection of mycoplasmas in host tissues are described in Chapter A6 in Vol. 1. The question of persistence and detectability of mycoplasmas during various stages of arthritis is of interest because of the inability to consistently and reproducibly isolate infectious agents from RA at any stage. In addition, determining the precise location of mycoplasmas within arthritic joints is important to an understanding of the pathogenic mechanisms of mycoplasmainduced joint damage. The M. arthriddis-rdbbit model may be particularly useful for studying detection methods because of the propensity of this organism to disappear from arthritic joints long before the inflammatory response abates. Methods used successfully in other animal models, as reviewed by Cole et al. (1985), include direct culture (in the case of M. arthritidis and M. pulmonis in rats and mice, active arthritis is associated with persistence of viable organisms in the joints) and antigen detection by enzyme immunoassay, immunoflourescence, and electron microscopy. Kohn and Chinookoswong (1989) used immunoperoxidase staining to localize M. pulmonis within rat joints. Earlier work by this group had indicated a preference for cartilage, particularly the lacunae, in neonatal animals; use of immunoperoxidase confirmed this for neonates and furthermore showed an age-related shift to synovial tissues in older animals. The efficacy of newer techniques, such as nucleic acid hybridization and polymerase chain reaction, has yet to be evaluated for the arthritis models, although DNA probes have been constructed for both M. arthritidis and M. pulmonis. Use of Models to Study Extraarticular Manifestations of Arthritic Diseases
Many of the experimental arthritides, particularly those of rats and mice, are actually systemic diseases, and while arthritis may be the most prominent manifestation, other organs are usually also involved. Similarly, the human arthritides are also often accompanied by signs and symptoms in other organs. Thirkill et aL (1992) use the M. arthritidis rat model to study arthritis-associated ocular inflammation. In one study, 6-week-old female Sprague-Dawley rats were injected with 1 X 10^ CFU M. arthritidis (ATCC 14152), and immune complexes containing mycoplasmal antigens were detected in circulation by antigen capture
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assays and Western immunoblotting and in ocular tissues by immunofluorescence. This may prove to be a useful model for elucidating mechanisms of the ocular inflammation that sometimes accompanies RA. Effect of Immunomodulation on Mycoplasmal Arthritides Athymic mouse models and various immunologic modulators in otherwise intact animals have been used to study immunologic aspects of mycoplasmal arthritides (Cole and Ward, 1979; Cole et al., 1985). Studies along this line have explored factors relating to disease persistence and specific and nonspecific aspects of resistance to infection. MECHANISMS OF PERSISTENCE
Binder et al. (1993) have studied the arthritogenicity of M. arthritidis for both young (47-81 days old) and adult (360 days old) Rowett nude rats. Arthritis was induced by an intravenous injection of 5 x lO^-lO'^ CFU of M. arthritidis strain ISRl. Infection was more severe and persisted longer in athymic rats than in their euthymic littermates and the adult nude rats were not protected against reinfection. The protective effect of antibodies against reinfection in immunologically intact animals is well known (Cole et al, 1985). This work demonstrates the importance of thymus-derived lymphocytes in recovery from M. arthritidis infection and in the development of a protective response. ROLE OF SPECIFIC IMMUNE RESPONSE IN PROTECTION
The inability of M. arthritidis to induce opsonizing and neutralizing antibodies in rats leaves open the question of which mycoplasmal antigens serve as targets of protective antibodies. It is now known that these antigens are present in a variety of M. arthritidis cell fractions (Washburn et al, 1992). We have shown that LEW rats could be protected by subcutaneous injection with a vaccine consisting of heat- or formalin-killed or sonically disrupted M. arthritidis (1 mg total protein) in incomplete Freund's adjuvant, followed by an intraperitoneal booster injection of 1 mg protein without adjuvant 3 weeks later. Rats could also be protected by vaccinating with mycoplasmal proteins sliced directly from SDS-polyacrylamide gels. In this experiment, gel fractions containing multiple antigens were used, but the technique should be applicable to individual proteins as well. Each dose consisted of mycoplasmal protein bands sliced from a single lane on which 200 jxg protein had been loaded. The first dose was emulsified in incomplete Freund's adjuvant and injected subcutaneously; additional injections were given intraperitoneally without adjuvant at weekly intervals. We gave five booster injections, but anti-M. arthritidis antibodies were detectable by ELISA within 1 week after the first boost. Rats were challenged by intravenous injection 1 week after the final boost with 1 x 10^ CFU of virulent M. arthritidis. Similar
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methods should prove useful in identifying a role in pathogenesis and protection for individual antigens and may be applicable to other species of arthritogenic mycoplasmas as well. ROLE OF NONSPECIFIC FACTORS IN RESISTANCE TO INFECTION
Although arthritis can be induced in rats and mice with relatively small numbers of organisms when the intraarticular route is used, induction by intravenous injection requires very large doses, indicating that nonspecific defenses must be very effective against systemic invasion with both M. arthritidis and M. pulmonis. This is supported by the fact that immunomodulators given prior to infection can enhance resistance of mice to M. pulmonis-'indxxctd arthritis (Cole et al, 1985). A macrophage-stimulating glycoprotein derived from Klebsiella pneumoniae was shown by Kaklamani et al. (1991b) to have a similar effect on resistance of mice to M. arthritidis. In the case of M. arthritidis in rats, avoidance of early, nonspecific host defenses appears to be an important virulence factor. However, even virulent strains are eventually cleared by rats, in which M. arthritidis tends to induce an acute self-limited infection, as opposed to mice, which are more prone to chronicity. A possible explanation for this difference comes from work by Kaklamani et al. (1991a), who showed that infection with M. arthritidis stimulated the phagocytic activity of fixed macrophages in both rats and mice but that this stimulation persisted longer in rats than in mice. They suggested that this might allow rats to effect a more rapid recovery. Use of Models to Study Immunologic Mechanisms of Joint Damage The immunology of many of the experimental mycoplasmal arthritides is complicated by the fact that the mycoplasmas themselves often exert profound immunomodulatory effects on the host animals. However, this ability is also of considerable interest when establishing a model for RA. Autoimmunity is one mechanism of joint tissue damage in RA, and biological mimicry between an arthritogenic agent and host tissues has been suggested as a possible mechanism for induction of autoimmunity (Cole et al., 1985). Following up on previous observations that anti-M. arthritidis (strain ISRl) antibodies in arthritic rats cross-reacted with normal rat tissues, Biische et al. (1990) showed that CD4+ T cell lines from an infected LEW rat also reacted with syngeneic chondrocytes, thus providing further evidence for a role for autoimmunity in the M. arthritidis rat model. Another possible mechanism of immunomodulation and induction of autoimmunity is under investigation at the University of Utah (see Chapter F7, Vol. I). All of the arthritogenic mycoplasmas exhibit some form of immunomodulatory activity, often in the form of nonspecific mitogenic stimulation of host lympho-
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cytes (Cole et aL, 1985). The most prominent and best understood of these systems is the M. arthritidis superantigen MAM. M. arthritidis is the only arthritogenic mycoplasma known to produce a superantigen, and its discovery has thrown a new light on M. arthritidis disease as a model for human arthritis. Although a precise role for MAM in rat and mouse arthritis remains undefined, it may participate by suppressing host responses and/or by activating T cells to release inflammatory cytokines. Purified MAM was shown to exert an inflammatory influence on synovial tissues on intraarticular injection into DA rats, in which it induced a significant, although short-lived, arthritic response (Cannon et aL, 1988). MAM injected into mice has been shown to induce T-cell anergy and B-cell activation in a way that may enhance development of autoimmunity. Exposure to MAM also has the ability to exacerbate type II collagen-induced arthritis in mice, a well-known autoimmune arthritis model. Also relevant to human disease is the discovery of an overlap between Vp usage by MAMreactive T cells and Vp usage in RA and other murine arthritis models. These activities and methods for the purification and characterization of MAM and other potential mycoplasmal superantigens are reviewed in Chapter F7 in Vol. I.
References Barden, J. A., and Tully, J. G. (1969). Experimental arthritis in mice with Mycoplasma pulmonis. J. Bacteriol. 100, 5-10. Binder, A., Gartner, K., Hedrich, H. J., Hermanns, W., Kirchhoff, H., and Wonigeit, K. (1990). Strain differences in sensitivity of rats to Mycoplasma arthritidis ISRl infection are under multiple gene control. Infect. Immun. 58, 1584-1590. Binder, A., Hedrich, H. J., Wonigeit, K., and Kirchhoff, H. (1993). The Mycoplasma arthritidis infection in congenitally athymic nude rats. J. Exp. Anim. Sci. 35, 177-185. Busche, K., Schleisier, M., Runge, M., Binder, A., and Kirchhoff, H. (1990). T-cell lines responding to Mycoplasma-arthritidis and chondrocytes in the Mycoplasma-arthritidis infection of rats. Immunobiology 181, 398-405. Cannon, G. W., Cole, B. C , Ward, J. R., Smith, J. L., and Eichwald, E. J. (1988). Arthritogenic effects of Mycoplasma arthritidis T cell mitogen in rats. J. Rheumatol. 15, 735-741. Cedillo, L., Constantino, G., Mayagoitia, G., Giono, S., Cuellar, Y., and Yanez, A. (1992). Experimental arthritis induced by Mycoplasma pneumoniae in rabbits. J. Rheumatol. 19, 344-347. Cole, B. C , and Ward, J. R. (1979). Mycoplasmas as arthritogenic agents. In "The Mycoplasmas" (J. G. Tully and R. F. Whitcomb, eds.). Vol. 2, pp. 367-398. Academic Press, New York. Cole, B.C., and Washburn, L. R. (1983). Evaluation of arthritogenic properties of mycoplasmas for small laboratory animals. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 305-310. Academic Press, New York. Cole, B. C , Ward, J. R., Jones, R. S., and Cahill, J. F. (1971). Chronic proliferative arthritis of mice induced by Mycoplasma arthritidis. I. Induction of disease and histopathological characteristics. Infect. Immun. 4, 344-355. Cole, B. C , Thorpe, R. N., Hassell, L. A., Washburn, L. R., and Ward, J. R. (1983). Toxicity but
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not arthritogenicity of Mycoplasma arthritidis for mice associates with the hapiotype expressed at the major histocompatibiUty complex. Infec. Immun. 41, 1010-1015. Cole, B. C , Washburn, L. R., and Taylor-Robinson, D. (1985). Mycoplasma-induced arthritis. In "The Mycoplasmas" (S. Razin and M. F. Barile, eds.). Vol. 4, pp. 108-160. Academic Press, New York. Cox, N. R., Davidson, M. K., Davis, J. K., Lindsey, J. R., and Cassell, G. H. (1988). Natural mycoplasmal infections in isolator-maintained LEW/Tru rats. Lab. Anim. Sci. 38, 381-388. Harwick, H. J., Kalmanson, G. M., Fox, M. A., and Guze, L. B. (1973). Arthritis in mice due to infection with Mycoplasma pulmonis. I. Clinical and microbiological features. J. Infect. Dis. 128, 533-540. Kaklamani, E., Karalis, D., Kaklamanis, P., Koumandaki, Y., Katsouyanni, K., Blackwell, C , Sparos, L., Weir, D., and Trichopoulos, D. (1991a). The effect oi Mycoplasma arthritidis infection on the phagocytic activity of macrophages in rats and mice. FEMS Microbiol. Lett. 76, 151-158. Kaklamani, E., Koumantaki, Y., Karalis, D., Rommain, M., Smets, P., Kaklamanis, P., Blackwell, C. C , and Weir, D. M. (1991b). Klebsiella pneumoniae glycoprotein RU-41741 enhances resistance of mice against Mycoplasma arthritidis-'mduced arthritis. FEMS Microbiol. Immunol. 76, 205-210. Kohn, D. F., and Chinookoswong, N. (1989). Detection of Mycoplasma pulmonis in arthritic joints of rats by indirect immunoperoxidase staining. Infect. Immun. 57, 1321-1323. Taylor, G., Taylor-Robinson, D., and Slavin, G. (1974). Effect of immunosuppression on arthritis in mice induced by Mycoplasma pulmonis. Ann. Rheum. Dis. 33, 376-384. Thirkill, C. E., Tyler, N. K., and Roth, A. M. (1992). Circulating and localized immune complexes in experimental mycoplasma-induced arthritis-associated ocular inflammation. Infect. Immun. 60, 401-405. Washburn, L. R., and Ward, J. R. (1988). Mycoplasma-induced arthritis in rabbits. In "Handbook of Animal Models for the Rheumatic Diseases" (R. A. Greenwald and H. S. Diamond, eds.), Vol. 1, pp. 109-124. CRC Press, Boca Raton, FL. Washburn, L. R., Hirsch, S., McKenzie, M., and Voelker, L. L. (1992). Vaccination of Lewis rats against Mycoplasma arthritidis-induccd arthritis. Am. J. Vet. Res. 53, 52-58.
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E5 EXPERIMENTAL INFECTIONS IN POULTRY Janet M. Bradbury and Sharon Levisohn
Introduction The term poultry encompasses domestic fowl, turkeys, ducks, geese, game birds, such as pheasant and partridge, and can also include pigeons. All these species have their own mycoplasma flora but extensive knowledge on the poultry mycoplasmas is restricted to the recognized pathogens, namely Mycoplasma gallisepticum and M. synoviae in the fowl and the turkey, M. meleagridis in the turkey, and M. iowae in the turkey embryo. The outcome of a mycoplasma infection, even with these known pathogens, can be profoundly affected by factors such as environmental stresses and the presence of other pathogens (Jordan, 1990a) and it is therefore essential to bear this in mind when studying experimental infections. For most purposes, birds used for experimental infections should be specific pathogen-free (SPF), or at least mycoplasma free, and at times it may be desirable to introduce another factor(s) in order for the pathogenicity of the mycoplasma to be expressed. When establishing an experimental infection the ultimate purpose will influence the procedures to be used. Areas frequently under study are those relating to pathogenicity, pathogenesis, and the immune response. Ideally these investigations require realistic reproduction of the natural disease, preferably induced by a natural route of infection. Under these conditions, there may be a great deal of bird-to-bird variation. Contact infection, which may be viewed as one of the more natural routes, can give very variable results. Areas of study such as evaluation of diagnostic methods and procedures for control necessitate a method 361 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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of infection which is relevant to the clinical situation of interest. Other common purposes include studies on efficacy of antimicrobials and protection by vaccines, in which cases a consistently reproducible challenge model is required, with results that can be quantified and subjected to statistical analysis. For this it may well be necessary to introduce the organisms in large numbers (> 10^ viable organisms) and by an unnatural route such as air sac or lung inoculation. Experimental infection of avian-derived material such as organ culture or embryonated eggs may be a convenient model system for studies of both avian and nonavian mycoplasmas. The production of specific antisera is still another area of interest, requiring different techniques. As well as the influence of environmental and concurrent infections referred to earlier, the outcome of mycoplasma infection in poultry is affected by factors such as the age and breed of the bird, and young birds will, in general, be more susceptible. The choice of organism is of paramount importance. Phenotypic changes may occur during multiple passages, and strains which have undergone multiple passage in vitro are likely to be attenuated. Thus the type strain may not be a suitable choice for many purposes, and some workers prefer to use a recent field isolate for experimental infection. Others have found that organisms reisolated from inoculated birds or embryos may show increased virulence. Different strains of the same Mycoplasma species may show a preference for one organ system over another. Even taking all this into account, it should be remembered that some strains of mycoplasma, even of the so-called pathogenic species, may have very low infectivity and virulence and may elicit an immune response only if administered intravenously. There are no generally accepted standard methods for producing experimental mycoplasma infections in poultry and the following are intended simply as guidelines. Much useful information on methods for producing experimental infections can be obtained by following up the references cited in the relevant chapters on the pathogenic mycoplasmas in Calnek et al. (1991).
Materials Inocula consist of mycoplasma broth cultures in logarithmic growth phase, containing a known number of viable organisms. It is desirable to use cloned cultures whose purity has been confirmed by a method such as immunofluorescence but, in order to keep in vitro passage level low, one filter cloning is recommended as a minimal purification step. The number of in vitro passages of the culture should be recorded. Inocula can be prepared in advance and frozen in aliquots and a viable count can be established on a thawed aliquot. A count
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should also always be conducted at the time of use. Relatively high numbers of organisms are often used in experimental infections (>10^). It should be noted that M. meleagridis does not normally grow to high titers in broth and it may be necessary to use centrifuged cultures or to wash the organisms off agar plates in order to produce a suitable inoculum. Host birds should be of defined status, e.g., SPF or mycoplasma free. Fertile eggs of the host species should also be derived from a mycoplasma-free flock since any of the pathogenic species can be transmitted through the Qgg. In addition, the presence of maternally derived antibodies can affect the outcome of experimental infection in ovo. Numbers of experimental animals should allow for adequate uninfected controls. Wing tags may be used if identification of individuals is needed. Bird husbandry should consist of standard procedures for working with infected animals. Limited access isolation housing with suitable ventilation, heating, food, and water should be used, or positive pressure isolator units when available. Brooding facilities are necessary for young birds. Precautions should be instituted to prevent dissemination of the infection, or transfer from one experimental group to another: for instance, a disinfectant bath should be supplied for dipping hands, feet, and any materials carried in or out of the poultry house (enclosed in polythene bags) and protective overalls and head covering, boots, and disposable gloves should be worn. A balance will be needed if weight gain and feed conversion are to be assessed. If embryo work is to be included, necessary equipment includes an egg incubator and egg candler.
Procedures Reproduction of Respiratory Disease ROUTE OF INOCULATION
Young birds (usually 1-14 days old) are inoculated via the nares or by eye drop using, for example, 0.05-0.1 ml of broth culture dispensed by an Eppendorf pipette. Alternatively the organisms can be inoculated directly into the infraorbital sinus or introduced into the trachea by means of a soft plastic cannula attached to a syringe. A microaliquoter with a disposable syringe (Scientific Manufacturing Institute) may be used to deliver a standard inoculum. Infection can be initiated by aerosol, which is usually administered for a standard time ( 5 10 minutes) to birds that are confined in a small closed area. The particle size influences the penetration of the inoculum, which in turn influences the course
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and outcome of infection. Reproducible results can be obtained using a standard medical nebulizer, such as DeVilbiss 45 (Somerset, PA) with controlled air pressure. For the purpose of assessing the protective effect of antimicrobials or vaccination, it is necessary to produce a quantifiable respiratory response, usually airsacculitis, with minimal bird-to-bird and group-to-group variation, usually the inoculum (0.05-0.2 ml, depending on the age of the bird) is introduced directly into the air sacs or lung of the young bird, using a very short narrow gauge hypodermic needle (Jordan, 1990b). Figure 1 indicates the correct location for lung inoculation. It is possible to determine the site of inoculation by introducing the same volume of saline with vital dye and immediately sacrificing an animal for examination of the relevant area. DOSE OF INOCULUM
A dose of >10^ viable organisms of a virulent strain of M. gallisepticum should be sufficient to produce clinical disease in turkey poults and possibly also in chicks, although they are less susceptible than turkeys. In chickens a superimposed infection with respiratory viruses, such as live infectious bronchitis or Newcastle disease virus vaccines, or with Escherichia coli will increase the
Fig. 1. Skeleton of the fowl indicating the site for direct lung inoculation. From Jordan (1990b).
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likelihood of M. gallisepticum causing overt respiratory disease. Some E, coli serotypes (e.g., 0 2 and 078) may act synergistically to give rise to colisepticemia with airsacculitis, pericarditis, and perihepatitis. Environmental factors such as increased ammonia levels or cold ) may increase the likelihood of respiratory signs and lesions in chickens. In the absence of some kind of exacerbating factors, M. synoviae will probably not give rise to respiratory signs in chickens or turkeys, although the latter may show sporadic sinusitis, and both species may develop airsacculitis. M. meleagridis and M. iowae may cause lesions in inoculated turkey air sacs but are unlikely to cause clinical signs. Details may be found in references quoted in the relevant chapters in Calnek et al. (1991). QUANTIFICATION
Respiratory disease is quantified by clinical and/or lesion scoring with additional criteria including isolation of the infecting agent. Serological responses can also be measured and in some cases may serve as an indicator of the efficacy of treatment or degree of protection. Respiratory signs such as coughing, sneezing, and tracheal rales can be noted by careful observation and listening, and nasal exudation is assessed for quantity and turbidity by squeezing the beak posterior to the nares. Ocular discharge and conjunctivitis are scored, as is swelling of the infraorbital sinuses. Lesions in dead birds are assessed macroscopically and/or microscopically, according to clearly stated criteria (see, for instance, Dykstra et al., 1985). Since mycoplasma infection may also result in reduced feed conversion and weight gain, these can be quantified and subjected to statistical analysis, if proper allowance is made for sex differences. Tracheal Explants and Organ Culture
Chick embryo tracheal organ cultures are sometimes used to assess the pathogenicity of respiratory mycoplasmas. Scoring systems are based on the vigor and/or extent of ciliary activity or cytopathologic changes (Levisohn et al., 1986; Dykstra et al., 1985). Tracheal rings are prepared by standard methods from chick embryos at 19 to 20 days of incubation. Mycoplasma inocula are prepared without thallium acetate as this could be toxic for the tracheal explants. Use of nonpermissive medium for organ culture and low inoculum levels will increase the likelihood that the course of disease in the model system reproduces that in vivo. Reproduction of Locomotory Disease
M. gallisepticum rarely produces locomotory disorders but M. synoviae introduced intravenously or into the footpad may give rise to synovitis and arthritis in
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the hock joint. Some strains may be more suitable than others for this purpose. Since the lesions of chronic M. synoviae arthritis seem to involve an immunopathological element, more satisfactory results are found using birds with greater immunological maturity (>2 weeks of age). Lesions may be quantified grossly by degree of swelling, exudation, and erosion in the joint and by microscopic changes (Morrow et al., 1990). Chondrodystrophy, accompanied by stunting and poor feathering, may be reproduced in turkeys by very early infection via the air sac route with M. meleagridis and some strains of M. iowae. Similar signs may be seen with M. iowae in chicks, particularly in heavier breeds (Bradbury and Kelly, 1991). Study of Egg Production and/or Egg Transmission and Infection of Reproductive System
Mechanisms of egg transmission are poorly understood. All the avian mycoplasma pathogens are transmitted through the egg to the next generation, but natural transmission is unpredictable and variable and hence very difficult to reproduce experimentally. For these reasons there is a paucity of information on this subject, particularly on the best methods to detect egg transmission. To simulate the effects of M. gallisepticum on egg production and transmission, female chickens are infected with a high dose of a virulent strain when at peak production (Glisson and Kleven, 1984). A combination of aerosol and intrasinus inoculation or air sac and sinus inoculation has been used. Sporadic egg transmission of M. synoviae can be obtained following the inoculation of laying hens by the respiratory or footpad routes, but it is unlikely to have a detectable effect on egg production. To evaluate the effects on egg production the percentage of production over a period is compared to that of sham-inoculated uninfected controls kept under similar conditions. The group numbers should be sufficient for statistical analysis. To evaluate rates of egg transmission and the effect of protective measures and treatment, isolation of mycoplasmas can be attempted from infertile eggs as well as from fertile eggs and dead-in-shell embryos. The yolk sac and vitelline membrane have both been found to harbor M. gallisepticum in experimental transmission studies. Little is known about the preferred sites for isolation of M. synoviae in infected eggs, although it has been cultured from infertile eggs and dead-inshell embryos and from the tracheas of day-old chicks. For M. meleagridis and M. iowae, which cause genital infections in the turkey, the condition can be simulated in the mature female by the introduction of contaminated semen. Resulting transmission of M. meleagridis may be detected by culturing from the shell membrane, vitelline membrane, or yolk and of M. iowae from the embryo gut or extraembryonic fluids. As with the other
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mycoplasma pathogens, the preferred sites have not been completely established and other sites may also yield the organism. Cytopathologic changes in the oviduct can be used to evaluate the protective effects of vaccine in the experimental infection of layer hens. Chicken or turkey oviduct explants have been suggested for use in a similar fashion to tracheal explants described earlier, although experimental conditions and suitability for such studies have not been well established. Production of Specific Antiserum Specific antisera can be prepared by parenteral inoculation of appropriate antigen into chickens, turkeys, or rabbits. Antigen for production of antisera in birds can be grown in artificial medium containing serum from the host species, with serum fraction, or in medium in which the serum has been replaced by liposomes. These measures are taken to avoid the production of unwanted antibodies to medium constituents. For birds the respiratory route of infection may be used and in this case the antibody response would be expected to mimic that found in the natural infection. Thus M. gallisepticum strains of known virulence and some M. synoviae strains can be inoculated via the respiratory tract to produce a humoral response, and the latter also may be introduced by the footpad route. Very young birds can be infected, but since they are not immunologically mature, a better antibody response will probably be obtained by inoculating 2- to 4-week-old birds or older. At least 2 weeks are needed before sampling. For less virulent strains, including most strains of M. synoviae, intravenous inoculation will probably give the most consistent results. M. meleagridis antibodies can be produced in mature chickens by intravenous inoculation, and young turkeys will produce antibodies following air sac inoculation of certain strains. Agglutinins to M. meleagridis have also been produced by oviduct inoculation of mature turkey hens (G. P. Wilding and M. Grant, personal communication). This is done by eversion of the oviduct, as practiced for artificial insemination, and transfer of organisms onto the mucosa by means of a cotton swab. M. iowae does not elicit a consistent humoral antibody response in birds infected by the respiratory, oral, or genital route. Antibodies have been produced in chickens following a series of intravenous inoculations. Rabbits can be hyperimmunized with all the recognized avian mollicute species by conventional methods (Senterfit, 1983). Such sera have good growth and metabolism-inhibiting properties, whereas antisera prepared in avians may not exhibit this property. However, adult chickens can be hyperimmunized by inoculation with antigen following a regime similar to that recommended for rabbits. The antigen may be presented with adjuvant, in which case certain oil-emulsion
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preparations (for instance, Titermax, CytRx Corporation, Technology Park, Atlanta, GA) will produce a good response with little tissue damage (S. H. Kleven, personal communication). Embryo Inoculation Embryos of any age can be inoculated, depending on the purpose of the experiment. For assessing pathogenicity, organisms are normally inoculated in 0.1-ml volumes into the yolk sac of 7- to 10-day-old chick embryos or 9- to 13day-old turkey embryos. For other avian species the equivalent stage of incubation is used. Mycoplasma inocula are prepared as described earlier but should be free of thallium acetate as it is toxic for embryos. Control eggs should be similarly inoculated with sterile medium. Eggs are incubated at 37°C, with turning, and are candled daily for viability. Deaths within 24 hours are discounted as nonspecific. Percentage embryo mortality and/or hatchability are assessed and embryos are examined for lesions and numbers of organisms. For some purposes, embryos may be inoculated with a low dose of mycoplasmas and the infected birds are allowed to hatch for the study of pathogenesis or treatment experiments. However, even very low doses of some mycoplasmas, especially M. iowae, may be lethal for the embryos. Procedures for Humane Killing of Birds and Embryos Birds are killed by intravenous or intraperitoneal injection of an overdose of a suitable anesthetic or by dislocation of the neck. Exposure to rising concentrations of CO2 gas is permitted in some countries for birds over 1 week of age and up to 1 kg body weight. Embryos can be killed by chilling at 4°C, by decapitation, or by injection of an overdose of a suitable anesthetic.
Discussion It is very important to remember that the mycoplasmas may be transmissible on the operator's clothing and hands after contact with infected birds and therefore adequate disinfection procedures are essential. Birds with overt respiratory signs may also generate aerosols containing mycoplasmas. Careful measures should be taken to protect any nearby poultry from risk of infection. Animal welfare is another important consideration in all experimental infections and should be taken into account when planning an experiment. In some countries strict procedures are laid down by law to ensure that minimum suffer-
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ing occurs. It is unusual for uncomplicated infections with avian mycoplasmas to cause severe clinical signs but birds that are in obvious discomfort should be killed humanely by an approved method, as indicated earlier. The complex nature of poultry mycoplasma disease was referred to in the introduction and it is not possible in this brief account to cover every possible aspect involved in producing experimental infections. As with other mycoplasmas, it may prove difficult to produce a consistently good result because the course of infection involves a complex and multifactorial relationship among mycoplasma, host, and environmental and other factors. Each laboratory will probably evolve its own favored methods but these may not necessarily be transferred successfully to another. An aspect of poultry mycoplasma infection that has not been addressed here is immunity. The immune system of birds differs from that of mammals in that the B cells are seeded in early life from the bursa of Fabricius, a discrete organ that can be excised by surgery or ablated by chemical treatment. Thus experimental mycoplasma infection in birds offers a unique opportunity for study of the role of the different arms of the immune system in defense and in immunopathogenesis. Techniques for both bursectomy and thymectomy are available in the relevant literature. Finally, it should be noted that poultry have the great advantage that the mycoplasma pathogens can be readily studied in their natural hosts, as well as in derivative model systems. Moreover, poultry are inexpensive to obtain and maintain in suitable numbers for statistical analysis of the results. Experimental infections with poultry mycoplasmas have a unique role in that they yield information which is of interest not only in comparative mycoplasmology but which can also provide answers to problems of epidemiology, diagnosis, and control of avian mycoplasmosis which is responsible for significant and worldwide economic losses in an important agricultural industry.
References Bradbury, J. M., and Kelly, D. (1991). Mycoplasma iowae infection in broiler-breeders. Avian Pathol. 20, 67-78. Calnek, B. W., Barnes, H. J., Beard, C. W., Reid, W. M., and Yoder, H. W., eds. (1991). "Diseases of Poultry," 9th ed. Iowa State Univ. Press, Ames. Dykstra, M. J., Levisohn, S., Fletcher, O. J., and Kleven, S. H. (1985). Evaluation of cytopathologic changes induced in chicken tracheal epithelium by Mycoplasma gallisepticum in vivo and in vitro. Am. J. Vet. Res. 46, 116-122. Glisson, J. R., and Kleven, S. H. (1984). Mycoplasma gallisepticum vaccination: Effects on egg transmission and egg production. Avian Dis. 28, 406-415. Jordan, F. T. W. (1990a). Avian mycoplasmoses. In "Poultry Disease" (F. T. W. Jordan, ed.), 3rd ed., pp. 74-85. Bailliere Tindall, London.
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Jordan, F. T. W. (1990b). Direct lung infection of chicks and turkey poults with mycoplasmas. Vet Rec. Ill, 502. Levisohn, S., Dykstra, M. J., Lin, M. Y., and Kleven, S. H. (1986). Comparison of m vivo and in vitro methods for pathogenicity evaluation for Mycoplasma gallisepticum in respiratory infection. Avian Pathol. 15, 233-246. Morrow, C. J., Bell, I. G., Walker, S. B., Markham, P. F., Thorp, B. H., and Whithear, K. G. (1990). Isolation of Mycoplasma synoviae from infectious synovitis of chickens AM^/. Vet. J. 61, 121-124. Senterfit, L. (1983). Preparation of antigens and antisera. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 401-404. Academic Press, New York.
E6 EXPERIMENTAL INFECTIONS OF SWINE Marylene Kobisch and Richard F. Ross
Introduction Mycoplasma hyopneumoniae is the primary agent of enzootic pneumonia, one of the most important chronic diseases in industrial pig herds (Ross, 1993). M. hyopneumoniae can cause experimentally induced pneumonia in pigs, and this model has been used to examine the pathogenesis of the infection (including attachment of the mycoplasma to target cells and pathogenetic mechanisms utilized by the microorganism in induction of pneumonia); clinical, microbiological, and serological aspects of the disease; and the efficacy of vaccines or antibiotics in the control of the infection. The various experimental models summarized in this chapter outline techniques useful to investigators working with pig mycoplasmas.
Materials M. hyopneumoniae Strains The strains used in these experiments are field strains (isolated from outbreaks of enzootic pneumonia). In the case of an old isolate, the frozen strain (-70° ) is cultivated in Friis liquid medium (1975) and is intranasally inoculated into one or two 2-week-old hysterectomy-produced piglets, reisolated from lung tissue, and then cultivated at 37° C in Friis medium. The strain is immediately used or is frozen until the experimental infection. After cultivation, the 371 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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organisms are harvested before the end of the exponential growth phase and are inoculated onto agar medium. Experimental Animals
Two types of piglets are used: piglets bom from Large White and Landrace sows obtained from a closed experimental hysterectomy-derived herd or hysterectomyproduced colostrum-deprived gnotobiotic piglets. The pregnant sows and the piglets are housed in separate units in which the air is forced through absolute filters to prevent any contact with infectious agents.
Procedure Experimental Infections and Laboratory Examinations
1. Naturally bom piglets are repetitively challenged at 2, 3, 4, and 5 days of age by the introduction into each nostril of 0.5 ml of a suspension of live M. hyopneumoniae organisms in Friis medium. The concentration of the organisms in the suspension is 10^ color-changing units (CCU) per ml. 2. Hysterectomy-produced colostmm-deprived gnotobiotic piglets are infected intratracheally by a sterile needle with 5 (for piglets 2 weeks of age) or 10 ml (for piglets 16 weeks of age) of broth cultures of M. hyopneumoniae (10^ CCU per ml). 3. An identical number of uninfected pigs receive broth medium using the same conditions. Rectal temperatures and clinical signs, notably coughing, are recorded daily. Blood samples are taken once a week from the piglets throughout the experiment and semm samples are analyzed by ELISA (see Chapter D8, this volume). At random intervals, from 2 to 22 weeks following inoculation, the piglets are anesthetized, euthanized by exanguination, and then necropsied. Postmortem observations are carried out on each animal and M. hyopneumoniae is detected by culturing from lung tissue and mucus from trachea (Friis, 1975). 4. Thin sections of fixed and paraffin-embedded lungs are stained with a mixture of hematoxylin and eosin and examined under a light microscope. For the detection of M. hyopneumoniae^ frozen thin sections of lungs are treated with anti-M. hyopneumoniae 3.ntibodiQS labeled with fluorescein isothiocyanate (Kobisch et al., 1978). Samples of trachea and bronchi from some piglets are collected and prepared as previously described for scanning electron microscopy (SEM) (Blanchard et ai, 1992). ELISA is performed as described by Nicolet et al. (1980).
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Discussion Clinical Symptoms
Age has no influence on the expression of the clinical signs. The body temperature of the control piglets is 39.5°C. In infected groups, a slight hyperthermia (40°-40.3°C) may be noticed 1 week after challenge. Coughing begins 2 weeks after infection, peaks after 5 weeks, and gradually declines. Twelve weeks postinfection, no clinical symptoms are noticeable. Growth is not retarded and appetites are normal. Postmortem Findings
No lesions are observed in the negative control group which received the broth medium. Necropsy observations show that lesions are confined to the lungs. Macroscopic lesions typical of mycoplasmal pneumonia are observed from 1 to 11 weeks postinoculation in pigs infected at 1 or 2 weeks of age and from 1 to 6 weeks in pigs infected at 16 weeks of age. These lesions are located in the ventral portions of the cranial and middle lobes, the accessory lobe, and the cranial portion of the caudal lobes of the lungs. In acute stages of disease, a catarrhal pneumonia is observed with exudate in the airways. The bronchial and mediastinal lymph nodes are often enlarged. In the chronic stages of disease, recovering lesions consisting of fissures of collapsed alveoli adjoining areas of alveolar emphysema are observed after 5 weeks in pigs infected at 16 weeks and after 9 weeks in pigs infected at 1 or 2 weeks of age. Histopathology
Pulmonary tissue modifications observed in the lungs of pigs in the acute phase of infection consist of hyperplasia of the epithelial cells (Fig. lA) and an increased perivascular and peribronchiolar accumulation of mononuclear cells. Lymphocytes and plasmocytes are both detected, but the latter are more abundant. As the disease progresses, perivascular- and peribronchiolarcharacteristic nodules, often compressing the lumen of the bronchioles, can be observed in recovering lesions (Fig. IB). Samples of trachea, collected from the infected piglets, show very mild lesions of tracheitis, consisting of epithelial hyperplasia and infiltration of the lamina propria by small numbers of lymphoplasmatic cells. The mucosal glands are normal. The lumen of the bronchi contains an exudate of mucoid material and polymorphous inflammatory cells with many neutrophils and some macrophages. In control pigs, lung tissue does not present any lesions.
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Fig. 1. The lung of a pig infected with M. hyopneumoniae showing (A) hyperplasia of the epithelial cells and (B) an increased perivascular and peribronchiolar accumulation of mononuclear cells. (C) SEM of a normal tracheal surface (C, ciliated cells; Ml, microvilli; M, mucus). (D) SEM of a pig trachea, 6 weeks postinfection with M. hyopneumoniae. The loss of cilia of epithelial cells and M. hyopneumoniae on the top of remaining cilia are observed.
Scanning Electron Microscopy
In the control pigs, normal tracheal and bronchial surfaces observed by SEM are composed of ciliated cells mixed with microvilli (Fig. IC). In the pigs killed
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at the first or second week after challenge, colonization of epithelial cells by M. hyopneumoniae is noticeable. Two to 8 weeks after inoculation, loss of cilia is evident, and in some cases, ciliated cells are seen to be exfoliating (Fig. ID). After this period, trachea and bronchi of pigs with recovering macroscopic lesions appear normal with some mucus production. In control pigs, the tracheal and the bronchial epithelium is ciliated. Serological Results
All inoculated pigs are seropositive 3 or 4 weeks after challenge. In the pigs infected at 1 or 2 weeks of age, the antibody response is similar in the earliest phase of the infection: peak titers are observed 10-12 weeks postinoculation and then decline regularly. All pigs are still seropositive 22 weeks after infection. In pigs infected at 16 weeks of age, ELISA titers are lower than in the two other groups, although antibodies are detected at the same time after infection. Recovery of M. hyopneumoniae
M. hyopneumoniae is isolated from the tracheal mucus and from the lung tissue at about 18 weeks after infection. The immunofluorescence test (Kobisch et al., 1978) performed on lung samples is positive for M. hyopneumoniae, generally in association with extensive pneumonia or in some cases with recovering lesions. A good correlation exists between these results and the isolation of M. hyopneumoniae which is generally not isolated from recovering lesions at the end of the experiment. At this time, the fluorescence test is negative. Data described earlier illustrate the utility of experimental models to study the effect of M. hyopneumoniae on the respiratory tract of piglets, although these models cannot completely reproduce the natural situation. These models are adequate and useful in evaluating the administration of medication in feed, in drinking water, or by injection of antibiotics, before or after the infection. Vaccines can be evaluated by the significant reduction in the severity of induced pneumonia or by the assessment of clinical signs in piglets passively or actively protected.
References Blanchard, B., Vena, M., Cavalier, A., Le Lannic, J., Gouranton, J., and Kobisch, M. (1992). Electron microscopic observation on the respiratory tract of SPF piglets inoculated with Mycoplasma hyopneumoniae. Vet. Microbioil. 30, 329-341. Friis, N. F. (1975). Some recommendations concerning primary isolation of Mycoplasma suipneumoniae and Mycoplasma flocculare: A survey. Nord. Veterinaer med. 27, 337-339.
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Kobisch, M., Tillon, J. P., and Vannier, P. (1978). Pneumonic enzootique a Mycoplasma hyopneumoniae chez le pore: Diagnostie rapide et reeherehe d'antieorps. Red. Med. Vet. 154, 847-852. Nieolet, J., Paroz, P., and Bruggman, S. (1980). Tween 20 soluble proteins of Mycoplasma hyopneumoniae as antigen for an enzyme-linked immunosorbent assay. Res. Vet. Sci. 29, 305309. Ross, R. F. (1993). Mycoplasmal diseases. In "Diseases of Swine" (A. D. Leman, B. E. Straw, W. L. Mengeling, S.D'Allaire, and D. J. Taylor, eds.), 7th ed., pp. 537-551. Iowa State Univ. Press, Ames.
E7 EXPERIMENTAL INFECTIONS IN CATTLE Ricardo F. Rosenbusch and H. Louise Ruhnke
Introduction The host specificity of molUcutes requires that experimental inoculations be performed in the host to understand the mechanisms of infection, the role of immune responses, and the etiological role that mycoplasmas may play in a specific disease. In planning experimental infections with bovine moUicutes, several considerations must be kept in mind. Cattle are hosts to many species of mollicutes, and several of these are pathogenic. Unless gnotobiotic calves are used to ensure freedom from mollicute infection, special conditions for raising mollicute-free calves have to be met or the cattle have to be proven to be free of the pathogen of interest by repeated isolation attempts and serology. Cattle are raised as outbred populations that are selected for specific production traits. As such, groups of calves in an experiment will exhibit significant variability of response to infections with mollicutes. Appropriate numbers of animals are needed, together with controls, to counteract this variability. Several infection models of current use in cattle are presented in this chapter. An ocular challenge model and several urogenital tract challenge models are described. These descriptions supplement those of respiratory tract challenge and intramammary inoculation which were included in Taylor (1983) and Howard (1983).
Ocular Challenge Cattle commonly harbor various species of mollicutes on their conjunctival mucosa. Infection with certain ubiquitous species {Mycoplasma bovoculi, 377 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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Ureaplasma diversum) occurs at birth, and other species of mollicutes infect cattle conjunctivae throughout their life. Additional species recovered in ocular conjunctivae of European and North American cattle are M. bovirhinis, M. arginini, Acholeplasma oculi, A. axanthum, A. laidlawii, M. dispar, M. bovis, M. bovigenitalium, and M. verecundum. Infectious conjunctivitis can be caused in calves by inoculation with M. bovoculi or U. diversum (Rosenbusch and Knudtson, 1980). In addition, cattle infected with M. bovoculi are more susceptible to ocular infections with gram-negative bacteria, including Moraxella bovis, which causes severe eye lesions associated with infectious bovine keratoconjunctivitis (Rosenbusch, 1983). The ocular challenge model in cattle can provide relevant insight into chronic mucosal infections by mycoplasmas at a site that can be easily and repeatedly sampled for organisms and immune responses. Cellular components of the immune response can also be sampled at a mucosal level (Norian and Rosenbusch, 1993). Calves for ocular challenge use are caught at birth into a sterile bag and transported immediately to an individual isolation facility. They are fed pasteurized colostrum or raised without colostrum. Calves are used for ocular challenge at 1 month of age or at more than 2 months of age if immunological maturity is needed. Calves should be shown to be free of mollicutes by repeated swab samplings from the lower conjunctival sacs and nasal passages. To prevent establishment of overabundant bacterial flora in the conjunctivas of calves, hay should be fed from raised feeders and the calves' tail mane should be shorn. Calves that have bacterial flora that may interfere with samplings (i.e., bacteria that overgrow in mycoplasma media) can be treated once with local antibiotics (bacitracin-neomycin-polymyxin veterinary ophthalmic ointment) 5 days prior to inoculation. Materials Calves, free of mollicute infections in conjunctival and nasal mucosa, and free of bacterial flora that may interfere with detection of mollicutes. Isolates of M. bovoculi should be cloned at least twice and used after low passage in modified Friis medium with 20% (v/v) fetal calf serum (Knudtson et al., 1986). Isolates of U. diversum should be grown in Hayflick's medium, pH 5.4, with 20% fetal calf serum. Media should be prepared without thallium acetate to avoid toxicity to host tissues. Syringes, 3 ml Sterile cotton-tipped swabs Sterile Kimura platinum spatula
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Slides, with multiple wells for cytology smears Sterile gauze pledgets, 2 x 2 cm and four layers thick Incubator at 3TC
Procedure INOCULATION
Calves are inoculated with a mycoplasma suspension of at minimum 10^ colony-forming units (CFU)/ml in growth media by instillation into each conjunctival sac. Three milliters of inoculum is added dropwise over a period of 3 minutes with gentle massage of the closed eyelids. SAMPLING
1. Samples for mycoplasma culture are conveniently taken with sterile cottontipped swabs which are rolled over the lower conjunctival sac. Swabs are transported in dry tubes on ice for prompt culture. 2. Sterile gauze pledgets can be placed in the lower conjunctival sac and left to collect fluid for 3 minutes. They are then removed and transported in dry tubes on ice. Tear fluid is expressed from the pledget by centrifugation of the pledget placed in a 3-ml syringe without plunger. Low-speed centrifugation (200 g) for 2 minutes at 25°C is sufficient to collect over 500 |JL1 of fluid from each eye into collection tubes attached to the syringe. For preservation of antibody activity, 1 mM EDTA is added prior to storing tear fluid samples at -20°C. 3. Exfoliative cytology samples are obtained by gentle curetting with a Kimura spatula. The cell and fluid-rich sample is deposited on a slide for histochemical or immunohistochemical processing. Semiquantitative estimation of cell populations in a sample can be obtained by counting cell types of interest and normalizing the count to 100 conjunctival epithelial cells.
Discussion
Sampling from the conjunctival sac induces mild and transient inflammation at the site. This reaction must be taken into account in interpreting data from colonization or immune response measurements. Swab sampling every 2 days and pledget sampling every 4 days do not appear to alter response parameters. Conjunctivitis can be assessed by clinical examination (ocular secretions, limbal vessel congestion) and by neutrophil responses detected in conjunctival exfoliative cytology samples. Pathogenicity of a mollicute for the bovine conjunctiva can be assessed by the
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duration of colonization and the titers of the agent recovered. With M. hovoculi and U. diversum, colonization exceeds 1 month in duration with sustained recoveries of 10^ or more organisms per swab (Rosenbusch and Knudtson, 1980).
Urogenital Tract Challenge Species of mycoplasmas that are most often isolated from the reproductive tract include U. diversum, M. bovigenitalium, M. canadense, and M. bovis. A. laidlawii is also a frequent isolate but thus far has not been shown to cause disease. Diseases associated with ureaplasmas, mycoplasmas, and acholeplasmas include granular vulvovaginitis, endometritis, salpingitis, abortion, and infertility in cows (Ball et al., 1987a,b; Doig et al., 1979; Kreplin et al., 1986; Miller et al., 1983; Stalheim and Proctor, 1976). The prepuce and semen are frequently colonized, and there may be loss of libido associated with some strains. Seminal vesiculitis has been reported and reproduced experimentally (Al-Aubaidi et al, 1972; Waelchli-Suter et al, 1982). Organisms may remain in the semen, even after it is treated with antibiotics and extended for artificial insemination, and subsequently infect cows during breeding. Embryos collected from infected animals following superovulation and insemination may also carry the organisms which are not removed by washing and thus carry the infection to the recipient. Young sexually mature heifers and bulls that have never been bred should be used for experimental inoculation of the reproductive tract. Young immature animals do not respond in the same way and give misleading results. Older mature animals may be more likely to have had previous exposure and may be resistant to new infections. Because gnotobiotic animals are not available for urogenital tract experiments, an equal number of matched control animals should be treated in the same manner as test animals to obtain meaningful and valid results. Controls should be inoculated before test animals and always sampled first after inoculation and at necropsy. Controls should be housed separately from test animals so that there is no contact between the groups. Materials Heifers should be free of abnormalities and should be cycling normally before use. Bulls should be producing normal spermatozoa. Mollicute isolates should be triple-cloned and at low passage (between 5 and 10 passages in artificial media). Sterile medium should be used to inoculate controls. Inoculate with 10^ to 10^ CPU in desired inoculum volume. Sterile cotton-tipped swabs
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Shielded insemination pipette for uterus inoculation Surgical equipment as required for amniotic sac inoculation or seminal vesicle inoculation
Procedures VULVA INOCULATION AND SAMPLING
1. Heifers should be negative when examined for moUicutes on preinoculation vulvar cultures taken on 6 different days over a period of at least 2 weeks. Heifers yielding pathogenic organisms (bacteria, viruses, and chlamydia) should not be used. 2. Prior to the day of inoculation the stage of the estrous cycle should be determined by rectal palpation. 3. Use inoculum of 10^ to 10^ mollicutes in 0.5 to 1 ml of media free of thallium acetate. 4. Gently wash the external vulva and dry with clean paper towel, then gently apply the culture to the vulvar mucosa using a sterile swab. Hold the vulvar lips closed for a few moments to allow the culture to be absorbed. 5. Postinoculation observations should include examination of the vulvar mucosa for hyperemia, edema, granularity, and discharge. Cows may show discomfort on examination when severe reaction is observed. UTERUS INOCULATION AND SAMPLING
1. Preinoculation, the same protocol as is used prior to vulvar inoculation to determine the absence of pathogens should be followed. It may be necessary to administer prostaglandin according to standard protocols to synchronize the cows so that they are all at the same stage in the cycle. 2. Inoculum of 10^ to 10^ organisms in 2 to 5 ml is used. 3. Thoroughly wash and dry the vulva and perineal region. To avoid contamination from the vulva, a standard shielded insemination pipette is guided through the cervix into the uterus by rectal manipulation. The inoculum is then deposited directly into the uterus. 4. If fertility is to be assessed, a single ejaculate from a proven bull should be processed, stored in liquid nitrogen, and used for all inseminations. Prior to use, 8 to 10 straws should be individually cultured to ensure the absence of mollicutes and pathogenic bacteria. The procedure to remove antibiotics prior to culture is given in Chapter D6 (this volume). There should be a delay of about 1 hour between insemination and inoculation to avoid possible inhibition of the organisms by the antibiotics in the semen.
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5. Postinoculation, clinical signs should be observed by examination of the vulvar mucosa daily for 10 days, then at 4-day intervals. Vulvovaginal swabs should be taken daily for 7 days, then weekly thereafter for microbiological examination. 6. Fecundity of the animals can be determined by serum progesterone concentration patterns by solid-phase radioimmunoassay 25 days postinoculation. 7. Antibody titers in cervical mucus may be of more significance than serum titers and may be determined for the isolate and for other pathogens of the reproductive tract. 8. The animals should be slaughtered at chosen intervals, and the reproductive tract removed intact and transported to the laboratory for collection of samples and culture or stored at -70°C within 4 to 6 hours. AMNIOTIC CAVITY INOCULATION
1. Pregnant cows should be kept isolated and cultured for mollicutes and pathogenic bacteria on three consecutive weekly cultures of the vulva. Serum should be examined for antibodies to known abortifacient infectious agents before and after the experiment. 2. Inoculum consists of 5 ml of a culture preparation containing a dose of 10^ to 10^ organisms. 3. The inoculum is introduced into the amniotic cavity by surgical procedure as described (Miller et al.y 1983). 4. Examine cows daily for the first week and then weekly until abortion or parturition. After delivery, samples of amnion and chorioallantois are cultured and examined microscopically. At necropsy of the fetus, samples of skin, lung, amnion, cotyledon, amniotic fluid, and stomach content are cultured for mollicutes and other known abortifacient infectious agents. Tissues are collected for histopathology. SEMINAL VESICLE INOCULATION OF BULLS
1. It is rare to find bulls that do not carry some mollicutes and bacteria, therefore bulls should be chosen that do not carry the organism to be used. Ideally, specific pathogen-free animals should be used, but are not available to everyone. Preinoculation cultures of the prepuce and semen should be made. Semen and seminourethral fluid can be obtained directly from the prolapsed penis by manual massage of the seminal vesicles, thus avoiding preputial contamination. By inoculating seminal vesicles it is possible to observe the spread of the infection in either ascending or descending direction. 2. Inoculum of 10^ to 10^ organisms in 2 to 5 ml is used. The culture is
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introduced into one seminal vesicle via the ischiorectal fossa (Waelchli-Suter et ai, 1982). An equal number of control bulls should be inoculated in a similar manner with sterile broth. 3. Monitor the infection by rectal examination, by examination of the semen for inflammatory cells, and by culture of the organism from seminourethral fluid. The bulls should be necropsied at chosen intervals and the complete urogenital tract examined by culture, immunology, and histopathology.
Discussion
After vulvar inoculation, very fine small raised granules, less than 1 mm in diameter, first appear in 3 to 5 days and are located lateral to the clitoris. Oblique light using a flashlight may be necessary to see the early granules. More marked granularity is readily evident and usually involves the lateral vulvar wall as well as the clitoral area. Discharge may be mucoid in mild cases or mucopurulent in more severe cases and may be seen only intermittently for a period of about a week. A classification of severe granular vulvitis is made when hyperemia, edema, granules, and discharge are present, whereas the absence of discharge is classified as moderate granular vulvovaginitis. Vulvitis will be observed within 3 weeks. If the organism is reisolated, associated with typical clinical signs, no other pathogens are recovered, and the control animals are negative on culture and clinical examination, an assessment of the virulence of the challenge organism may be made. Uterine pathogenicity of an inoculated organism can be assessed by correlating clinical signs, reisolation of the organism, gross and histopathologic lesions, and cervical mucosa antibody levels (if a test is available). After amniotic cavity inoculation, placentitis may be observed but there may be no gross lesions in the fetus. Microscopic lesions may be seen in various tissues of the fetus depending on the mollicute inoculated (Miller et al., 1983; Stalheim and Proctor, 1976). Although inoculation of the seminal vesicle is highly artificial, it may allow some conclusions as to the pathogenicity of a mollicute strain if there is an increase in sperm abnormalities accompanied by inflammatory cells and if the organism is isolated from the gland and/or other sections of the tract and there is a specific rise in antibody titers.
References Al-Aubaidi, J. M., McEntee, K., and Lein, D. H. (1972). Bovine seminal vesiculitis and epididymitis caused by Mycoplasma bovigenitalium. Cornell Vet. 62, 581-596. Ball, H. J., Armstrong, D., McCaughey, W. J., and Kennedy, S. (1987a). Experimental intrauterine inoculation of cows at oestrus with Mycoplasma canadense. Vet. Rec. 120, 370.
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Ball, H. J., Armstrong, D., McCaughey, W. J., and Kennedy, S. (1987b). Experimental intrauterine inoculation of cows at oestrus with a bovine ureaplasma strain. Ir. Vet. J. 41, 371-372. Doig, P. A., Ruhnke, H. L., and Palmer, N. C. (1979). Experimental bovine genital ureaplasmosis. II. Granular vulvitis, endometritis and salpingitis following uterine inoculation. Can. J. Comp. Med. 44, 259-266. Howard, C. J. (1983). Intramammary inoculation of cattle and other animals. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 2, pp. 299-304. Academic Press, New York. Knudtson, W. U., Reed, D. E., and Daniels, G. (1986). Identification of mycoplasmatales in pneumonic calf lungs. Vet. Microbiol. 11, 79-91. Kreplin, C. M. A., Ruhnke, H. L., Miller, R. B., and Doig, P. A. (1986). The effect of intrauterine inoculation with Ureaplasma diversum on bovine fertility. Can. J. Vet. Res. 51, 440-443. Miller, R. B., Ruhnke, H. L., Doig, P. A., Poitras, B. J., and Palmer, N. C. (1983). The effects of Ureaplasma diversum inoculated into the amniotic cavity in cows. Theriogenology 20, 367374. Norian, L., and Rosenbusch, R. (1993). Mycoplasma Z7c>vocM//-augmented bovine natural killer activity. Comp. Immunol. Microbiol. Infect. Dis. 16, 113-122. Rosenbusch, R. (1983). Influence of mycoplasma preinfection on the expression of Moraxella bovis pathogenicity. Am. J. Vet. Res. 44, 1621-1624. Rosenbusch, R., and Knudtson, W. (1980). Bovine mycoplasmal conjunctivitis: Experimental reproduction and characterization of the disease. Cornell Vet. 70, 307-320. Stalheim, O. H. V., and Proctor, S. J. (1976). Experimentally induced bovine abortion with Mycoplasma agalactiae subsp. bovis. Am. J. Vet. Res. 37, 879-883. Taylor, G. (1983). Respiratory challenge of experimental animals. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 2, pp. 291-298. Academic Press, New York. Waelchli-Suter, R. O., Doig, P. A., Ruhnke, H. L., Palmer, N. C , and Barker, C. A. V. (1982). Experimental genital ureaplasmosis in the bull. Schweiz. Arch. Tierheilkd. 124, 273-295.
E8 EXPERIMENTAL INFECTIONS OF PLANTS BY SPIROPLASMAS X. Foissac^ J. L. Danet, C. Saillard^ R. F. Whitcomb, and J. M. Bove
Introduction Although many spiroplasmas are associated with insects, only three spiroplasma species (Spiroplasma citri, S. kunkelii, and S. phoeniceum) inhabit the phloem of infected plants and are phytopathogenic (Calavan and Bove, 1989; Whitcomb, 1981) S. citri, the causal agent of citrus stubborn disease, and 5. kunkelii, the causal agent of com stunt disease, are transmitted by leafhoppers (Homoptera: Cicadellidae) that imbibe sap from sieve tubes. S. phoeniceum was discovered after natural transmission to periwinkle (Catharanthus roseus). The natural vector of this agent is unknown, but the spiroplasma has been transmitted to aster by an unusual vector, Macrosteles quadrilineatus, after microinjection of cultured spiroplasmas. The life cycle of S. citri after acquisition by the insect host begins with multiplication in the intestinal epithelium, followed by passage through the gut barrier (Liu et al., 1983b). The spiroplasma then spreads throughout the hemolymph and colonizes many tissues, including the salivary glands. After passing through the salivary cells, helical spiroplasma cells enter the salivary duct and are injected into the phloem of plants during feeding. Because spiroplasmas have never been transmitted to plants mechanically, experimental infection must be accomplished by using leafhopper vectors to inoculate plants. Three procedures can be used for experimental infection of the insect: 385 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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(i) acquisition achieved by exposing noninfected leafhoppers to infected plants, (ii) acquisition as a result of feeding through Parafilm membranes, and (iii) microinjection. For the three known plant spiroplasmas, all of which can be readily cultivated, the two last methods are especially convenient (Markham and Townsend, 1979; Markham and Oldfield, 1983). This chapter presents an example of the technique's application: experimental infection of Circulifer haematoceps, the vector of citrus stubborn disease in the Mediterranean area (Fos et al., 1986) by S. citri, and transmission to periwinkles. In all tests, females are preferred to males or last instar nymphs because their longevity is greater under experimental conditions. (For experimental phytoplasma plant infections see Chapter E9 in this volume.)
Materials For Insect Microinjection Periwinkle (Ca. roseus) seedlings, 5-7 cm tall Noninfected Ci. haematoceps reared on Mathiola incana at 28° , with a photoperiod of 17 hours Cages and insect-handling equipment (aspirator) Microinjection apparatus (Markham and Townsend, 1979) Binocular microscope Rubber tubing Peristaltic pump Needle/tubing connector made from glass tubing Hand-drawn needles. Pyrex tubing is heated in a hot gas flame and drawn to approximately one-fourth of its original diameter. The capillary needles are obtained by drawing the narrowed tube in a cool gas flame to an approximate diameter of 10-20 |xm. Needles are autoclaved before use. Carbon dioxide source Fine forceps Frozen surface (an ice block wrapped in Parafilm M) to keep the insect anesthetized or a multiple insect holder (Markham and Townsend, 1979)
For Membrane Feeding Feeding apparatus (Fig. 1) Acquisition buffer: 20 mM phosphate buffer, pH 7.2; 5% (w/v) sucrose
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upper parafilm membrane glass bead (diameter 2-3 mm) lower parafilm membrane spiroplasma culture nylon screen plastic vial window 50mm
hole with a cork nylon screen moistened cotton 20mm
Fig. 1 . Apparatus for membrane feeding. Insects are introduced into the apparatus by the hole closed by a cork. Moistened cotton and a window are used for humidification and ventilation.
For S. citri Culture and Determination of the Infection Rate in Insects SP-4 liquid and solid medium (1% agarose) for spiroplasma culture (Whitcomb, 1983) Insect-grinding apparatus 5-ml syringe Syringe filter (pore diameter 0.45 ixm)
Procedures Microinjection Female leafhoppers are anesthetized by exposure to CO2 for a few seconds, then restrained dorsal side down on a frozen surface, immobilized on an insect holder, or maintained in a continuous flow of moist CO2. The needle is filled with about 30 |JL1 of a mildly acidic spiroplasma culture (10^ CFU/ml). The insects are then microinjected with about 0.1 |xl of inoculum between two of the last abdominal stemites; this injected volume is sufficient to cause a visible swelling of the abdomen. The injected leafhoppers are caged on healthy stock plants in the greenhouse at 30°C for an incubation period of 12 days. Membrane Feeding The surfaces of the Parafilm membranes used in the apparatus are sterilized by immersion in 70% (v/v) ethanol for a few minutes before use. Leafhoppers are
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placed in the feeding apparatus and starved for 20 hours. The S. citri cells in culture are concentrated 10-fold by centrifugation (14,000 g, 20 minutes, 20°C) and the pellet is resuspended in acquisition buffer. Then a 400-|xl sample of acquisition buffer, containing the concentrated 5. citri culture (5 x 10^ colony forming units (CFU/ml), is placed between the two membranes. After feeding for 24 hours at 20°-25°C, leafhoppers are transferred onto healthy stock plants in the greenhouse at 30°C, for an incubation period of at least 14 days. Determination of Insect Infection Rate by S. citri The spiroplasma titer in CPUs can be determined by anesthetizing groups of one to five leafhoppers and crushing them in 2 ml of SP-4 medium in insectgrinding tubes. The extracts are passed through a syringe and filter (pore diameter 0.45 |jLm). Tenfold serial dilutions are made in SP-4 medium, and 0.2 ml of each dilution is plated on SP-4 plates. Plates are incubated at 32°C. Transmission to Periwinkles After completion of the incubation period, leafhoppers are caged on small periwinkles, individually or in groups of 5 to 10 insects. The transmission period is at least 2 days for infected leafhoppers. Leafhoppers infected by membrane feeding are caged on periwinkles until they die. After transmission, periwinkles are treated by insecticide to kill surviving insects and are kept in the greenhouse at 30°C. Typical symptoms appear after 3-4 weeks (Calavan and Oldfield, 1979).
Discussion The infection rates of Ci. haematoceps by S. citri and rates of transmission to periwinkles by infected insects are dependent on the method of infection. After microinjection, all insects carry about 10^-10^ CPU after injection and 10^-10^ CPU at the end of the incubation period. All exposed insects are infected and transmit to healthy plants with 20-30% efficiency. The transmission rate varies among 5. citri strains. Some strains are transmitted poorly or well while other strains are not transmitted. Strains that have been passed repeatedly in culture tend to lose their ability to complete the biological cycle. Because microinjection does not allow study of the passage of S. citri through the insect gut barrier, membrane feeding must be employed for some experiments. Immediately after feeding, S. citri can be isolated from 50 to 80% of insects fed through membranes; titers range from 10^ to 10^ CPU per insect. In contrast, only 3-10% of insects are infected in 24 hours after aquisition feeding. After 14 days of incuba-
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tion on healthy stock plants, only about 50% of the infected adults are able to transmit infection (1.5% efficiency). Similar experiments have been performed by Liu et al. (1983a) with last nymphal stages of Ci. tenellus, one of the American vectors of S. citri, and by Rana et al. (1975) with other leafhoppers that transmit S. citri. Acquisition feeding and microinjection, but not membrane feeding, have been used for studies (Whitcomb, 1989) on Dalbulus maidis and com, to which the com stunt spiroplasma, S. kunkelii, is readily transmitted. Acquisition feeding from infected com is especially efficacious with this agent; high rates of infection can be obtained by feeding early instar nymphs on diseased com plants or on detached leaves from diseased plants in moist chambers. Spiroplasma pathogenicity in insect hosts can also be investigated using microinjection and/or membrane acquisition procedures (Clark and Whitcomb, 1983).
References Calavan, E. C , and Bove, J. M. (1989). Ecology of Spiroplasma citri. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.), Vol. 5, pp. 425-485. Academic Press, San Diego, CA. Calavan, E. C , and Oldfield, G. N. (1979). Symptomatology of spiroplasmal plant diseases. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.). Vol. 3, pp. 37-64. Academic Press, New York. Clark, T. B., and Whitcomb, R. F. (1983). Special procedures for demonstration of mycoplasma pathogenicity in insects. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 69-79. Academic Press, New York. Fos, A., Bove, J. M., Ali, Y., Brun, P., and Vogel, R. (1986). La cicadelle Neoaliturus haematoceps (Mulsant & Rey) est vecteur de Spiroplasma citri en Mediterranee. Ann. Inst. Pasteur/Microbiol. 137A, 97-107. Liu, H. Y., Gumpf, D. J., Oldfield, G. N., and Calavan, E. C. (1983a). Tranmission of Spiroplasma citri by Circulifer tenellus. Phytopathology 12t, 582-585. Liu, H. Y., Gumpf, D. J., Oldfield, G. N., and Calavan, E. C. (1983b). The relationship of Spiroplasma citri and Circulifer tenellus. Phytopathology 73, 585-590. Markham, P. G., and Oldfield, G. N. (1983). Transmission techniques with vectors of plant and insect mycoplasmas and spiroplasmas. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 261-267. Academic Press, New York. Markham, P. G., and Townsend, R. (1979). Experimental vectors of spiroplasmas. In "Leafhoppers Vectors and Plant Disease Agents" (K. Maramorosch and K. F. Harris, eds.), pp. 413-445. Academic Press, New York. Rana, G. L., Kaloostian, G. H., Oldfield, G. N., Granett, A. L., Calavan, E. C , Pierce, H. D., Lee, L M., and Gumpf, D. J. (1975). Acquisition oi Spiroplasma citri through membranes by homopterous insects. Phytopathology 65, 1143-1145. Whitcomb, R. F. (1981). The biology of spiroplasmas. Annu.lRev. Entomol. 26, 397-425. Whitcomb, R. F. (1983). Culture media for spiroplasmas. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, pp. 147-158. Academic Press, New York. Whitcomb, R. F. (1989). Spiroplasma kunkelii: Biology and ecology. In "The Mycoplasmas" (R. F. Whitcomb and J. G. Tully, eds.). Vol. 5, pp. 487-544. Academic Press, San Diego, CA.
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E9 EXPERIMENTAL PHYTOPLASMA INFECTIONS IN PLANTS AND INSECTS Alexander H. Purcell
General Introduction Insect vector transmission is central to studies of plant pathogenic moUicutes. All such plant pathogens either have known insect vectors or have epidemiological data strongly suggesting that a mobile vector is responsible for natural spread of the diseases. For plant-pathogenic or other insect-associated mollicutes, experimental infection is required to study the transmission process or the pathology of the mollicutes to insects. The first requirement for such studies is that the host insect can be reared in the laboratory, preferably free of all other plant or insect pathogenic mollicutes. Each species of insect has optimal rearing conditions in the laboratory; appropriate literature references for a specific insect or a species with similar requirements should be consulted for guidance in rearing methods. Protocols for transmission of mollicutes by insect vectors have been reviewed (Whitcomb, 1972a,b; Markham, 1982; Markham and Oldfield, 1983). Methods for evaluating mycoplasma pathogenicity to insects have been reviewed by Whitcomb et al. (1983) and Clark and Whitcomb (1983). This chapter presents simple, general protocols for insect transmission of plant pathogenic phytoplasmas following (a) feeding insects on an infected plant, (b) injecting insects with suspensions of mollicutes using glass needles, and (3) feeding insects through a membrane on a solution in which mollicutes are suspended. (See also Chapter E8, this volume, for experimental spiroplasma infections in plants and insects.) 391 Molecular and Diagnosti'- Procedures in Mycoplasmology, Vol. II
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Feeding Insects on a Phytopiasma-Infected Plant Materials One hundred or more young, pathogen-free adult or late-instar nymphs of aster Icafhopptv Macrosteles quadrilineatus or other Macrosteles species capable of transmitting the aster yellows phytopiasma organism (AYO). A symptomatic aster (Callistephus chinensis) or common plantago (Plantago major) plant infected with AYO and an identical healthy aster or plantago plant (negative control). A simple plastic tube aspirator (25-40 cm long x 0.4-0.5 cm inside diameter with a 3-cm-diameter circle of fme Dacron mesh secured to one end by snugly fitted flexible plastic or rubber tubing about 35-55 cm long). Two insect-proof cages large enough to contain the entire phytopiasma source plant or cages that fit tightly over the plants to prevent escape of insects. The simplest cage is a sac of fine mesh Dacron organdy cloth that contains the entire plant with the mouth of the sac tightly secured at the base of the plant with string or rubber bands. Cylindrical plastic cages to enclose test plants. Healthy aster or plantago plants 1-2 weeks after transplanting into 6- to 11-cmdiameter plastic nursery pots. Aerosol container of acephate or other systemic insecticide suitable for spraying directly onto test plants.
Procedure 1. Transfer about half of the healthy insects from the rearing cage onto the AY-diseased source plant and place the other half onto the healthy control plant using the aspirator. Keep the plants at 22°-26°C for 7-14 days, then aspirate individual or groups of leafhoppers onto the test plants enclosed in a small cage. 2. After 1 week, transfer insects from the original test plants to new test plants, using an aspirator. After removing insects, the plants should be sprayed with insecticide and kept in a glasshouse in light and temperature conditions suitable for good plant growth, i.e., 22°-26°C. 3. If transmission efficiency data are desired, repeat step 2 for as long as desired. 4. Symptoms of AY should appear within 3-8 weeks in the glasshouse. Diagnostic serological, DNA hybridization, or polymerase chain reaction (PCR) assays for the AY organism can be used to confirm phytopiasma colonization of test plants.
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Discussion
Temperature may significantly affect phytoplasma transmission, the time required for leafhoppers to become infectious (latent period), the time required for symptoms to appear in plants (incubation period), and the severity of symptoms. The probability of successful acquisition or infection by leafhoppers increases rapidly with access time on the source plant or test plant, respectively, but maximum efficiency for both is usually approached within a few days to a week (Purcell, 1982; Chiykowski, 1973). These times can be varied experimentally to estimate their effects on transmission efficiency. If the objective is to simply produce plants infected by AYO, late-stage (fourth to fifth instar) leafhopper nymphs can be placed on infected plants for 3-4 weeks, then transferred in groups (10-50 each) to larger susceptible healthy plants. Older plants generally are less susceptible to infection with AYO than are young plants (Chiykowski, 1973), so that larger numbers (10-20) of infective leafhoppers are needed to ensure a high percentage of vector transmission to older plants. These general methods for feeding, acquisition, and inoculation can be adapted to other mollicute plant pathogens, including spiroplasmas (see Chapter E8, this volume), and to other vector species.
Microinjection of Phytoplasma Insect Vectors Microinjection can be used to infect insect vectors with mollicutes from culture (to prove Koch's postulates, for example), to determine the infectivity of phytoplasmas purified from insects or plants, to study mechanisms involved in vector transmission, and to increase transmission efficiency by vectors (Whitcomb, 1972b; Purcell, 1979) or by insects that normally do not transmit after feeding on infected plants (Markham and Townsend, 1979). The general protocol presented here for the aster yellows phytoplasma uses the simplest possible methods and materials but has been used successfully for numerous other mollicute plant pathogens. Materials
All materials used in preceding protocol One hundred or more pathogen-free aster leafhoppers Tissue grinder, 1-2 ml Sterile 0.45-|jLm filter for filter sterilizing small quantities (<1 ml) of fluid Syringe to fit into sterile filter or filter holder Fine forceps
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Binocular dissecting microscope Crushed wet ice in bowl or insulated bucket Microcentrifuge Sterile 1.5-ml plastic centrifuge tubes Sterile plastic petri dishes (100-mm diameter) Carbon dioxide gas (with regulator to reduce pressure) or frozen (dry ice) fragments covered with a 2- to 4-cm layer of cotton and placed in a plastic container (400-800 ml) with a snap-on cap or Parafilm Flask (500 ml) containing 200 ml water with an inlet tubing that passes through a rubber stopper to bubble CO2 gas below the water level and an outlet tube in the same stopper. Healthy aster or plantago plants large enough to support 100 leafhoppers for 1-3 weeks and small aster or plantago test plants Plant labels Glass needles for injection (See Discussion for further details) with rubber tubing (25-40 cm long) that fits securely over the base end of the needle Plastic 100-mm-diameter petri dish bottoms filled with water and covered with a smooth layer of thinly stretched Parafilm and frozen in a horizontal position.
Procedure 1. Position cages over healthy plants on which the injected insects will be placed during their latent period (2-3 weeks). Label plants. 2. Collect healthy leafhoppers into a small, transparent holding cage about 30-60 minutes before injections are to begin. 3. Prepare inoculum of the phytoplasma to be transmitted. Macerate 100 live aster leafhoppers that are infective with AYO in 1 ml of 0.3 A^ glycine, 0.3 A^ sucrose (or sucrose added to total osmolarity of 800 mosmols), adjusted to pH 7.0 with NaOH or HCl. Centrifuge for 1 minute at 1000 g. Filter supernatant through a sterile 0.45-|jLm membrane filter into a sterile 1.5-ml tube. Maintain inoculum on crushed ice and use within 2 hours. 4. Pipette separate drops (10-20 |JL1) of inoculum into a sterile petri dish by spotting each drop separately at the bottom of the dish. 5. Prepare needle by loading it into needle holder and suction tube. Load needle with liquid inoculum while viewing under a dissecting microscope and gently sucking until a small amount of liquid is visible in needle. 6. Anesthetize leafhoppers with carbon dioxide gas humidified by bubbling through water in a flask. First, aspirate the leafhoppers into a small, freely ventilated holding cage. Then place the cage into a metal or plastic can, fill can with CO2 gas, and seal the top of can with a snug lid or membrane for 30-40 seconds. An alternative is to put fragments of frozen CO2 into the bottom of a
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plastic cup, insulate the dry ice with a 2-cm layer of cotton, and seal the cup with a lid. The CO2 is heavier than air and will remain at the bottom of the container. A small cage with the leafhoppers can be placed in the cup to anesthetize insects. 7. Gently sprinkle the anesthetized insects into a large petri dish lid placed under a dissecting microscope. By gently shaking the dish, most of the insects will come to rest on their backs. As an option, the insects can be placed instead on a membrane-covered, wet ice-filled petri dish so that the ice will keep the insects cool and inactive. 8. Position the forceps in one hand to prevent the leafhopper from sliding away during injection, and with the other hand gently jab the needle through the intersegmental membrane of one of the abdominal segments (second to sixth) with the needle pointed toward the insect's head. A slight, rapid jab may be needed to penetrate the membrane but be careful not to penetrate the body wall too deeply and pierce internal organs. Gently blow out inoculum into leafhopper to produce a barely visible swelling of the abdomen. If the legs of the insect begin to extend, too much inoculum is being injected. Immediately remove injected insects to a small holding cage. They will usually remain upright but inactive for several minutes after regaining consciousness. 9. Transfer injected insects to caged plants. 10. Repeat steps 4-8 with dilutions of phytoplasma inoculum or with buffer for negative controls. 11. Transfer insects to new test plants individually or in groups, as desired. After removing insects, the plants should be sprayed with insecticide and kept in a glasshouse in light and temperature conditions suitable for good plant growth. 12. Symptoms of AY should appear within 3-8 weeks in the glasshouse.
Discussion
Glass microinjection needles can now be obtained commercially. The simplest injection needles can be made by hand over a small flame from the tips of Pasteur pipettes or 2- to 4-mm-diameter flint glass tubing. The first pull is made by manually centering and attempting to stretch a length of flint glass tubing (3-4 mm diameter) over a small gas flame until the glass begins to melt and stretch. At that point, the glass should be simultaneously removed from the flame and rapidly stretched to the maximum extent possible without breaking. A second pull of the narrowest but still hollow portion of the stretched glass tubing is made in the same manner over a small flame of an alcohol burner. The needles should be packed into a glass petri dish on top of a layer of cotton or sealed in a culture tube, sterilized by autoclaving, and dried aseptically in a warm oven or incubator before use. For injection the needle should be securely inserted into a section (25-35 cm) of flexible tubing so that inoculum can be drawn up by
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suction and expelled by gentle pressure using mouth aspiration or other devices (Markham, 1982). If mouth suction is used, a filter apparatus (e.g., syringefitting filters) should be connected between the needle and the operator's mouth. Practice is needled to become efficient at injecting insects, and this should first be done with buffer inoculum. Only anesthetize and inject small groups (5-10) of insects before anesthetizing additional insects as some of the insects will regain consciousness and become active, and longer or repeated periods of exposure to CO2 will kill many of the anesthetized insects. A more rapid method of injecting large numbers of insects is to place the anesthetized insects onto a shallow, flat-faced circular piston, over which is fitted a cylindrical sleeve that is covered at one end with thinly stretched Parafilm. Lowering the sleeve over the piston immobilizes the insects under the thin film of Parafilm; the insects are then injected through the film (Caudwell, 1977). This method has the advantage of inoculating many insects in a short time, but it is more difficult to precisely inject each insect in the abdomen. Needles should be changed after every 10-20 insects or more often if the needle becomes dull. Mortality from trauma during injection usually is evident within the first day after the injection.
Membrane Feeding Materials Noninfective leafhoppers to be fed through membranes Small cylinders (1- to 3-cm diameter, 2-4 cm long) cut from clear flexible or rigid tubing Corks or rubber stoppers to fit snugly into one end of the cylinders Parafilm cut into 3- to 8-cm squares 70% ethanol Inoculum of phytoplasma from in vitro cultures concentrated by centrifugation (> 10,000 g for 10-20 minutes) and resuspended in 0.5 to 0.9 M sucrose, pH7.0 Sterile micropipettes and pipetter Sterile laminar flow hood
Procedure 1. Sterilize Parafilm M squares by immersion in 70% ethanol for 2 minutes followed by drying under a sterile air flow.
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2. Perform steps 2-5 in a sterile laminar flow hood to maintain sterility in the inoculum fed to insects. Stretch a sterile Parafilm M square lengthwise and then 90° in the other direction to the maximum extent without breaking the film. 3. Fasten the stretched membrane over the open end of a small (1- to 3-cm diameter) cylinder so that it is smooth and taut. 4. Pipette inoculum onto membrane until drop covers about 50-70% of the surface within the inside diameter of the cylinder. 5. Lower a freshly stretched square of Parafilm gently over the drop and tilt and press the membrane to one edge of the cylinder, then carefully continue to stretch and lower the membrane so as to encircle the drop and seal the entire outer edge of the upper membrane to the lower membrane. Tamp the upper outside edge of the cylinder to ensure that the two membranes are sealed completely around the drop. With practice this can be done so that all air is excluded from around the drop of fluid. 6. Gently introduce an insect into the cylinder with an aspirator and plug the bottom of the cylinder with a cork or stopper. 7. Allow the insects to feed through the membrane on the inoculum for several hours or overnight. 8. Remove the insects and place them on test plants as in the preceding two protocols. 9. Diagnosis of plant disease symptoms in test plants should be made and recorded as in the previous two protocols.
Discussion
Phytoplasmas have not been successfully transmitted by feeding through a membrane on liquid inoculum, but not many such attempts have been reported. Spiroplasma kunkelii has been successfully transmitted by the com leafhopper Dalbulus maidis using a similar protocol (Chen and Liao, 1975). Membrane feeding often is much less efficient than either natural feeding acquisition or injection of vectors. Innumerable variations of containers to hold insects during membrane feeding can be adapted to this basic procedure. For example, inoculum can be introduced into vials and insects fed on the opposite side within a cylindrical cage that fits onto the vial or the inoculum can be pipetted into wells in a multiple-well plastic tray covered with a single large stretched membrane, and insects are confined individually over each well in small cylindrical cages (e.g., 1.5-ml plastic centrifuge tubes). Yellow light above the feeding insects reportedly improves feeding by homopterous insects (Koyama, 1973). Various types and concentrations of sugars, minerals, and lipids can improve the amount of feeding on artificial diets (Alivizatos, 1982).
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References Alivizatos, A. S. (1982). Feeding behavior of the spiroplasma vectors Dalbulus maidis and Euscelidius variegatus in vivo and in vitro. Ann. Inst. Phytopathol. Benaki 13, 128-144. Caudwell, A. (1977). Un appareil permetant d'immobiliser les insectes a injecter dans les epreuves d'infectivite des jaunisses. Ann. Phytopathol. 9, 521-523. Chen, T. A., and Liao, C. H. (1975). Com stunt spiroplasma: Isolation, cultivation, and proof of pathogenicity. Science 188, 1015-1017. Chiykowski, L. N. (1973). Factors affecting the infection of plants with clover phyllody agent transmitted by Macrosteles fascifrons. Ann. Entomol. Soc. Am. 66, 987-990. Clark, T. B., and Whitcomb, R. F. (1983). Special procedures for demonstration of mycoplasma pathogenicity in insects. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 369-379. Academic Press, New York. Koyama, K. (1973). Preference for color of diets in Inazuma dorsalis (HemipteraiDeltocephalidae). Jpn. J. Appl. Entomol. Zool. 17, 49-53. Markham, P. G. (1982). The 'yellows' plant diseases: Plant hosts and their interactions with the pathogens. In '"Plant and Insect Mycoplasma Techniques" (M. J. Daniels and P. G. Markham, eds.), pp. 82-100. Halsted Press, New York. Markham, P. G., and Oldfield, G. N. (1983). Transmission techniques with vectors of plant and insect mycoplasmas. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 261-267. Academic Press, New York. Markham, P. G., and Townsend, R. (1979). Experimental vectors of spiroplasmas. In "Leafhopper Vectors and Plant Disease Agents" (K. Maramorosch and K. F. Harris, eds.), pp. 413-445. Academic Press, New York. Purcell, A. H. (1979). Transmission of the X-disease agent by the leafhoppers Scaphytopius nitridus and Acinopterus angulatus. Plant Dis. Rep. 63, 549-552. Purcell, A. H. (1982). Insect vector relationships with procaryotic plant pathogens. Annu. Rev. Phytopathol. 397-417. Whitcomb, R. F. (1972a). Bioassay of clover wound tumor virus and the mycoplasmalike organisms of peach western X and aster yellows. V.S., Dep. Agric, Tech. Bull. 1438, 1-32. Whitcomb, R. F. (1972b). Transmission of viruses and mycoplasma by Auchenorrhynchous homoptera. In "Principles and Techniques in Plant Virology" (C. I. Kado and K. Agrawal, eds.), pp. 168-203. Van Nostrand-Reinhold, New York. Whitcomb, R. F., Clark, T. B., and Vaughn, J. L. (1983). Pathogenicity of mycoplasmas for arthropods and its possible significance in biological control. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 361-367. Academic Press, New York.
E10 MYCOPLASMAS AND IN VITRO INFECTIONS OF CELL CULTURES WITH HIV Shyh-Ching Lo and Alain Blanchard
Introduction Some species of mycoplasmas have been found with a significantly higher frequency among patients with AIDS (acquired immunodeficiency syndrome) (Dawson et al., 1993; Wang et al., 1992, 1993). In vitro studies have shown that mycoplasmas can markedly enhance the cytopathogenicity of HIV-1 (human immunodeficiency virus type-1). Specifically, infections of the normally lowvirulence budding human retrovirus in cultures of CD4+lymphocytes produce prominent cytocidal effects in the presence of some species of mycoplasmas and/or their biological products (Chowdhury et al., 1990; Lo et al., 1991; Lemaitre et al., 1992). The observations suggest that these AIDS-associated mycoplasmas may play an important role in the pathogenesis of AIDS. To better understand the interaction between the mycoplasmas and HIV-1, as well as their molecular mechanisms in pathogenesis, further studies are needed. However, cell culture conditions, distinct growth phases of the infected cells in cultures, and the infectious units of both mycoplasmas and HIV-1 introduced into cultures may all significantly affect the ultimate outcomes in cell biology studies. This chapter attempts to define the experimental conditions which, in our experience, produce more consistent and more meaningful data with in vivo implications. Mycoplasmas such as Mycoplasma fermentans, M. genitalium, M. pirum, or M. penetrans apparently do not require the presence of a specific receptor mole399 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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cule on the mammalian cell surface to grow in cell cultures. However, generally, the retrovirus HIV-1 can only infect and replicate in cells with positive surface CD4 molecules. Thus, CD4+ human lymphocyte cell lines (CEM, HUT-78, H9, Molt-3, etc.), monocytic cell lines (U937, THPl), or phytohemagglutinin (PHA)stimulated human peripheral blood mononuclear cells (PBMC) have been the most commonly used cell cultures in studies of HIV-1. It is important to note that the sudden introduction of any mycoplasma alone at a high titer into cell cultures may produce significant cytopathic effects. Thus, in cell biology studies, cultures should be infected with lower titers of infectious mycoplasmas that normally do not produce significant cytopathic changes in cell cultures. Another possibility is to establish a chronic mycoplasmal infection by cultivating the cells in the presence of mycoplasmas for 2-3 weeks until the cytopathic effects disappear. This chapter describes the specific conditions for human lymphocyte culture and methods to measure HIV-1 or mycoplasmal replication in cell cultures following different periods of infection.
Materials and Methods
Cell Cultures 1. Logarithmically growing lymphocytes (5 x lO^) in culture of 25-50 ml RPMI 1640 medium supplemented with 10% fetal calf serum (Gibco, Grand Island, NY) are infected with 1 x 10^ HIV-1 infectious units (see Assay of Infectious HIV Virions) and 1 x 10^ mycoplasma infectious units. 2. Cells are incubated at 37°C for 2 hours and are then washed once with fresh culture medium to remove nonadherent viral and mycoplasmal particles. 3. Changes of cell number in cultures are determined every day after trypan blue exclusion. In order to account for variations in the kinetics of HIV infection, it is necessary to perform these assays in duplicate or even triplicate.
Measurement of HIV Replication Normally, one of the most sensitive ways to detect retroviruses is to assay their unique enzyme activity reverse transcriptase (RT). However, many mycoplasmas produce a potent inhibitor(s) that affects the RT enzyme assay (Quillent et al., 1995; Vasudevachari et al., 1990). The inhibition is most likely due to mycoplasmal endonucleases which rapidly destroy the cDNA product in the assay.
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Thus, the measurement of HIV repUcation in cultures containing mycoplasmas may have to rely on the measurement of HIV-specific antigens p24/p25 or the infectious virions in the infected cultures. ELISA FOR HIV-1-SPECIFIC p24/p25 ANTIGENS
1. One hundred microliters of culture supernatant is saved every day for the quantitative testing of the virus-specific antigens. 2. The procedures of ELISA are performed in strict accordance to the instruction of the commercial kit (Coulter Corp., Hialeah, FL). 3. Using an ELISA reader, an absorbance reading is recorded for each well in the plates at 410 nm. ASSAY OF INFECTIOUS HIV VIRIONS
1. An additional 100 |xl of supernatant is also obtained from the cultures every day for the measurement of titers of infectious HIV-1 virions. 2. The culture supernatant is serially diluted 10-fold (up to lO^^), and 100 |JL1 is inoculated into fresh cultures of CEM (or other CD4+ lymphocyte) cell lines. 3. The cultures inoculated with serially diluted supernatant containing various concentration of infectious virions are incubated for another 10 to 12 days. 4. The supernatant from each of these cultures is tested for the presence of HIV-1-specific p24/p25 antigens. The titers of infectious HIV particles in the original culture supernatant can be determined in this way.
Measurement of Mycoplasma Proliferation
1. Another 100 |xl of supernatant is also withdrawn from the cultures every day for the measurement of mycoplasmal titers. 2. The culture supernatant is serially 10-fold diluted (up to lO^^) in SP4 broth media (see Chapter A2 in Vol. I) or another appropriate medium adequate for the optimal growth conditions of the mycoplasma under study. Each 5 ml of SP4 culture media is inoculated with 0.1 ml of supernatant dilution. 3. The mycoplasmal cultures inoculated with the serially diluted supernatant containing various concentration of viable mycoplasmal organisms are incubated at 3TC for 1 week. 4. Acidic changes of the SP4 medium indicate mycoplasmal growth. The titers of infectious HIV particles in the original culture supernatant can be determined in this manner.
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Cytopathic Effects in Lymphocytes Syncytia formation in lymphocyte cultures induced by HIV and its actual cell killing effects were found to be separate events (Garry, 1989, Lo et ai, 1991). To document the cytocidal effects of HIV-1 in CD4+ lymphocytes, the viable cells in cultures, with or without mycoplasmas, are counted daily using a hemacytometer after trypan blue exclusion. To minimize sampling errors, the cells are counted routinely in triplicate. Cytokine Production in HIV- and Mycoplasma-lnfected Cultures Various cytokines are known to be produced by lymphocytes and macrophages following HIV-1 infection. It is not surprising that various mycoplasmas may also induce the production of cytokines in cultures of human lymphocytes and macrophages, with or without HIV infection (Miihlradt et al., 1994). It is still not clear if a cytokine(s) plays a significant role in the HIV cell killing effects of lymphocytes. Specific assays measuring various cytokines are discussed in other sections (see Chapters F5 and F6 in Vol. I).
Discussion It is necessary to make sure that cell cultures as well as viral stocks used in cell biology studies are free of contamination by mycoplasmas. The most sensitive assays that are available need to be used to verify that the HIV and cell culture stocks are mycoplasma free and are described in Section F of this volume. Studies of infection by a particular virus can unknowingly be dealing with a mixed infection of both virus and mycoplasma(s). In many instances, the biological effect identified in cell culture studies as due to a particular virus was actually produced by occult infections by contaminating mycoplasmas. Indeed, many commonly used HIV-1 viral stocks, previously propagated in various human lymphocyte cultures, have been contaminated by different cell culture mycoplasmas such asM. arginini, M. hominis, M.fermentans, and Acholeplasma laidlawii. In order to demonstrate that the effects are actually due to the presence of mycoplasmas, it is possible to perform the same experiments in the presence of antibiotics (minocycline, pefloxacin, and others) (Lemaitre et al., 1992). In this case, it is necessary to include the proper control showing that these antibiotic compounds do not have a cytotoxic effect by themselves. The deliberate use of mycoplasma-contaminated cell cultures also represents a great risk of contamination for the other cell cultures maintained in the laboratory. The most stringent care should be taken to prevent this problem (McGarrity et al., 1983) (see Chapter F6 this volume).
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Since both HIV-1 and AIDS-associated mycoplasmas are considered potentially biohazardous, laboratory practices handling these infectious agents should strictly follow biosafety level 2 (B2) precautions. Detailed guidelines of B2 practice are listed in Biosafety Microbiological and Biomedical Laboratories, HHS Publication No. (CDC) 88-8395.
References Chowdhury, M. I. H., Munakata, T., Koyanagi, Y., Kobayshi, S., Aral, S., and Yamamoto, N. (1990). Mycoplasma can enhance HIV replication in vitro: A possible co-factor responsible for the progression of AIDS. Biochem. Biophys. Res. Commun. 3, 1365-1370. Dawson, M. S., Hayes, M. M., Wang, R. Y.-H., Armstrong, D., Budzko, D. B., Kundsin, R. B., and Lo, S.-C. (1993). Detection and isolation of Mycoplasma fermentans from urine of HIV positive patients with AIDS. Arch. Pathol. Lab. Med. Ill, 511-514. Garry, R. F. (1989). Potential mechanisms for the cytopathic properties of HIV. AIDS 3, 683-694. Lemaitre, M., Henin, Y., Destouesse, F., Ferrieux, C , Montagnier, L., and Blanchard, A. (1992). Role of mycoplasma infection in the cytopathic effect induced by human immunodeficiency virus type 1 in infected cell lines. Infect. Immun. 60, 742-748. Lo, S.-C, Tsai, S., Benish, J. R., Shih, J. W.-K., Wear, D. J., and Wong, D. M. (1991). Enhancement of HIV-1 cytocidal effects in CD4+ lymphocytes by the AIDS-associated mycoplasma. Science 251, 1074-1076. McGarrity, G. J., Gamon, L., and Sarama, J. (1983). Prevention and control of mycoplasmal infection in cell cultures. In "Methods in Mycoplasmology" (J. G. TuUy and S. Razin, eds.) Vol. 2, p. 203-208. Academic Press, New York. Quillent, C , Grau, O., Clavel, F., Montagnier, L., and Blanchard, A. (1994). Inhibition of HIV-1 reverse transcriptase assay by nucleases produced by contaminating mycoplasmas. AIDS Res. Hum. Retroviruses, 10, 1251-1257. Vasudevachari, M. B., Mast, T. C , and Salzman, N. P. (1990). Suppression of HIV-1 reverse transcriptase activity by mycoplasma contamination of cell cultures. AIDS Res. Hum. Retroviruses 6, 411-416. Wang, R. Y.-H., Shih, J. W.-K., Grandinetti, T., Pierce, P. F., Hayes, M. M., Wear, D. J., Alter, H. J., and Lo, S.-C. (1992). High frequency of antibodies to Mycoplasma penetrans in HIVinfected patients. Lancet 340, 1312-1316. Wang, R. Y.-H., Shih, J. W.-K., Weiss, S. H., Grandinetti, T., Pierce, P. F., Lange, M., Alter, H. J., Wear, D. J., Davies, C. L., Mayur, R. K., and Lo, S.-C. (1993). Mycoplasma penetrans infection in male homosexuals with AIDS: High seroprevalence and association with Kaposi's Sarcoma. Clin. Infect. Dis. 17, 724-729.
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SECTION
F
Diagnosis of Mycoplasma Infections of Cell Cultures
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F1 INTRODUCTORY REMARKS Joseph G. Tully
Although the first report of cell culture contamination with mollicutes (mycoplasmas) was made in the mid-1950s (Robinson et al, 1956), the continuing worldwide occurrence of these organisms in numerous continuous and primary cell lines, in hybridomas, and in a variety of other in vitro cell systems and their products remains a serious and formidable problem. More importantly, this contamination represents a significant factor in compromised or invalidated research findings and in enormous economic losses in research time, materials, and industrial production. Without adequate recognition by more cell culturists of the importance of mycoplasma contamination and effective detection schemes, solutions to this situation will remain elusive. Some national and international regulatory requirements have been established for the monitoring and testing for mycoplasmas in cell lines and seed stocks used in the production of therapeutic products destined for both humans and animals. However, few countries have developed testing programs or monitor the increasing application of cell culture systems in both public and private clinical diagnostic laboratories. Numerous efforts have been made over the intervening years to define the pervasive effects of mycoplasma infections on the eukaryotic cell. Today, it still remains no overstatement to say that every parameter of cell structure, growth, and metabolism can be affected by this infection. The extent of these changes has been well detailed over the years and in a number of reviews (McGarrity and Kotani, 1985; McGarrity et al, 1992; Barile and Rottem, 1993). As stated elsewhere (McGarrity and Kotani, 1985), the reported incidence of mycoplasma infection of cells can be very misleading. Infection depends on the type of cells examined, methods of detection, laboratory practices, and a number 407 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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of other conditions. However, there is some general impression that probably 10-15% of cell lines examined in the United States and in Europe are infected, with perhaps higher incidences in some other geographic locations. The identification of particular Mycoplasma or Acholeplasma species in cell infections can be a useful exercise as it may give some clue as to the origin of the infection. Bovine or other animal mycoplasmas or acholeplasmas in cell infections frequently originate in the serum supplements (i.e., fetal bovine), whereas mycoplasmas of human origin usually can be traced either to contamination of the primary specimen (i.e., lymphocytes, macrophages, etc.) or to breaks in sterility within the cell culture laboratory. Animal mycoplasmas, particularly M. arginini andM. hyorhinis, continue to be some of the most frequent species identified in cell infections. This remains puzzling since it had been thought that significant improvements had occurred in the quality assurance programs of most commercial sources of animal sera. Presently, most serum is usually obtained within a closed collection system that avoids much of the earlier contamination with filterable organisms, and some commercial laboratories have introduced techniques that subject final serum lots to filtration through membranes with pore diameters as small as 35 nm. However, animal serum should still be viewed as a critical factor in any cell culture control program. Not all commercial lots of serum are tested for mycoplasmas by the most appropriate methodology or are subjected to other than a 450-nm filtration, an exercise which fails to remove the organisms from the product. Of course, breaks in quality control tests and even in 35-nm filtration can also occur. Evidence exists that the number of M. fermentans strains isolated from cell infections has significantly increased since the mid-1980s, although few attempts to document this change have been recorded (McGarrity et al., 1992). Since this organism has been identified in human primary blood specimens (see Chapter D5, this volume), it is a common belief that this increase relates to the increased application of primary human cells in culture. Human-derived lymphocytes and macrophages are now frequently employed in cell culture applications in both immunology and in viral research (such as AIDS). Likewise, M. pirum, which appeared to be a cell contaminant with no well-established host or origin, has now been identified in several primary human lymphocytic cell cultures (see Chapter D5, this volume). However, this species seems to occur much less frequently than M. fermentans as a contaminant. Unfortunately, the occurrence of other human mycoplasmas in cell culture systems (especially M. ovale and M. salivarium) is still being recorded, but in many instances in numbers less than M. fermentans. These species are commensals in the oropharynx of humans and are thought to enter cell cultures through aerosols from the respiratory tract or via mouth pipetting. In Section F, the contributions selected continue to emphasize the important role that multidetection schemes play in the diagnosis of mycoplasma infections
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of cells. The experiences obtained since the 1970s have amply confirmed the value of a methodology that involves cultivation attempts on artificial media (broth and agar) and inoculation of a clean indicator cell line. The indicator cells, grown on coverslips and then inoculated with test cell material, are stained separately with a DNA stain and a conjugated antiserum to M. hyorhinis (see later). Culture methods with proper quality controls on the medium formulation employed are still a crucial aspect of detection, and several variations in a suggested scheme are outlined in Chapter F2. Culture techniques should always be evaluated on how well they correlate with the results of the indicator cell system. The authors have also provided an approach to the cultivation of M. hyorhinis cultivar a strains (Gardella and Del Giudice, 1995). These organisms, all of which have been identified in cell culture infections, are inhibited by a variety of standard mycoplasma medium components. The indicator cell system, with a conjugated antiserum specific for these organisms, allows their detection in the absence of axenic cultural documentation. Other techniques suggested for detection of such M. hyorhinis strains (Kotani et al., 1990) have not been confirmed in other laboratories. The use of a clean indicator cell system in conjunction with DNA stain and immunofluorescence was first developed by Del Giudice and Hopps (1978), and was intended to provide detection of M. hyorhinis strains that were incapable of growing on standard mycoplasma media. In Chapter F3, a method is described that utilizes staining by both techniques on a single coverslip, a procedure updated from an earlier report (Freiberg and Masover, 1990). The polymerase chain reaction (PCR) for the detection of cell culture mycoplasmas has been exploited rather extensively during the past few years. However, some of the reports have been difficult to evaluate since PCR results have not been correlated with results of conventional broth and agar methods or with indicator cell methodology. Since the PCR technique will also amplify DNA from both viable and nonviable mycoplasmas, one must also consider whether the PCR results should be confirmed in other testing protocols that measure viability of suspected contaminants. Chapter F4 outlines a PCR technique that has been successfully applied in a large-volume cell culture laboratory. The recommended approach to the treatment of mycoplasma infection in cell lines is to discard the cells rapidly, evaluate whether other cell lines in the laboratory are infected, determine the possible source of the infection, and then start with new cells from a reputable source. However, when the cell line or hybridoma line is of unquestioned value, elimination of mycoplasmas by antibiotic treatment should be considered. This should only be done under special circumstances as the procedure is time-consuming and quite labor-intensive. In Chapter F5, the authors have evaluated some of the earlier approaches to mycoplasma eradication from cells and have outlined a technique that they have
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successfully performed under a wide range of circumstances. The key to success also requires that the follow-up period after treatment allows for a number of cell passages in antibiotic-free media and that an effective detection method be used to reassess the possible reemergence of mycoplasmas in the treated cell line. Finally, chapter F6 covers some of the important areas in the prevention and control of mycoplasma infections in the cell culture laboratory. As is emphasized, the continued use of antibiotics in the normal maintenance of cell lines is undoubtedly one of the most critical factors responsible for the problem. Antibiotics mask the cell to cell spread of infection and significantly block an appropriate signal to the investigator that a breakdown in technique has occurred. In addition, the presence of antibiotics in cell cultures used for monitoring contamination has a direct and adverse effect on both detection by culture techniques and on the results obtained in the indicator cell test.
References Barile, M. F., and Rottem, S. (1993). Mycoplasmas in cell culture. In "Rapid Diagnosis of Mycoplasmas" (I. Kahane and A. Adoni, eds.), pp. 155-193. Plenum, New York. Del Giudice, R. A., and Hopps, H. E. (1978). Microbiological methods and fluorescence microscopy for the direct demonstration of mycoplasma infection on cell cultures. In "Mycoplasma Infection of Cell Cultures" (G. J. McGarrity, D. G. Murphy, and W. W. Nichols, eds.), pp. 57-69. Plenum, New York. Freiberg, E. F., and Masover, G. K. (1990). Mycoplasma detection in cell culture by concomitant use of bisbenzamide and fluoresceinated antibody. In Vitro Cell Dev. Biol. 26, 585-588. Gardella, R. S., and Del Giudice, R. A. (1995). Growth oi Mycoplasma hyorhinis cultivar a on semisynthetic medium. Appl. Environ. Microbiol. 61, 1976-1979. Kotani, H., Butler, G. H., Tallarida, D., Cody, C , and McGarrity, G. J. (1990). Microbiological cultivation oi Mycoplasma hyrohinis from cell cultures. In Vitro Cell Dev. Biol. 26, 91-96. McGarrity, G. J., and Kotani, H. (1985). Cell culture mycoplasmas. In "The Mycoplasmas" (S. Razin and M. F. Barile, eds.). Vol. 4, pp. 353-390. Academic Press, New York. McGarrity, G. J., Kotani, H., and Butler, G. H. (1992). Mycoplasmas and tissue culture cells. In "Mycoplasmas: Molecular Biology and Pathogenesis" (J. Maniloff, R. N. McElhaney, L. R. Finch, and J. B. Baseman, eds.), pp. 445-454. Am. Soc. Microbiol., Washington, DC. Robinson, L. B., Wichelhausen, R. H., and Roizman, B. (1956). Contamination of human cell cultures by pleuropneumonia-like organisms. Science 124, 1147-1148.
F2 ISOLATION OF MYCOPLASMAS FROM CELL CULTURES BY AXENIC CULTIVATION TECHNIQUES Richard A. Del Giudice and Joseph G. Tully
General Introduction The demonstration of characteristic colonial growth on axenic solid medium is a sensitive and specific method to detect mycoplasma infection of cell cultures and it is the standard against which all new methods must be gauged. This is true of all mollicutes known to infect cell cultures, with the exception of Mycoplasma hyorhinis cultivar a strains (Del Giudice et al., 1980). The fact that M. hyorhinis cultivar a strains do not grow on conventional mycoplasma media makes it necessary to use an indicator cell culture system to detect their presence (Del Giudice and Hopps, 1978; Chapter F3, this volume). Cultivar a strains have been grown on axenic solid medium (Gardella and Del Giudice, 1995), but this medium has not yet been tested for direct isolation from infected cells. Thus indirect methods are still required to detect cultivar a strains (Del Giudice and Gardella, 1984; McGarrity and Barile, 1983; McGarrity and Kotani, 1985). For all other mollicutes found in cell cultures, cultivation on solid medium is more sensitive than indicator cell cultures (Del Giudice and Gardella, 1984). This chapter presents culture media formulations and liquid and solid medium techniques that have been developed since the 1960s. These techniques and the indicator cell culture system have been integral parts of a complete mycoplasma test and this combined approach to mycoplasma isolation is highly recommended. 411 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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Materials for SP-4 Broth and Agar Preparation Frozen tissue culture cell suspensions (at -70°C) for testing SP-4 mycoplasma broth and agar medium (see Chapter A2, Vol. I, for details of medium preparation and quality controls) Gaspak incubation system (BBL Microbiology Systems, Cockeysville, MD), using envelopes for either anaerobic or carbon dioxide environments Sterile, glass, screw-cap culture tubes (15 x 100 mm) Sterile, plastic culture plates (60-mm diameter) Sterile, 1-dram screw-cap glass vials Standard microscope, with 100-200x magnification Laminar flow hood
Procedure 1. In the laminar flow hood, thaw the frozen cell suspension containing both cells and supernatant fluid and add about 1.0 ml of the suspension to a screw-cap glass culture tube containing about 9 ml of SP-4 broth. Tighten the cap, identify the cell line on the tube, and incubate the tube at 37°C. Always include an uninoculated tube of the current lot of medium for comparative and control purposes. 2. At the same time, inoculate each of two SP-4 agar plats with 0.2 ml of the cell suspension. Allow the suspensions to dry on the agar medium while in the hood. Again, identify the cell line on the plate. Place one agar plate each in the Gaspak jar for either anaerobic incubation or incubation in carbon dioxide environment (37°C). Immediately refreeze the cell suspension at -70°C for possible use in indicator cell lines or for repeat culture tests. 3. Observe the SP-4 broth tube at daily intervals and subculture when the culture exhibits turbidity or pH change. Subculturing should include plating of 0.2 ml to each of two SP-4 agar plates, and inoculation of 0.2 ml to a 1-dram vial containing 2 ml of SP-4 broth. A shift to a lower pH in the initial SP-4 broth tube is not necessarily presumptive of mycoplasma growth since metabolic activity of the cells in the broth medium can lower the pH. 4. Examine microscopically (160-200 x) the two agar plates inoculated with the primary cell suspension after incubation for 3-5 days and again after 7-14 days. Compare plates incubated in either anaerobic or carbon dioxide environments for growth. Agar colonies will usually appear in 3-7 days, and more frequently on plates incubated anaerobically. Mycoplasmas might not appear as fried egg-like colonies on initial growth, and it is necessary to distinguish colonies from individual tissue cells or clumps of cellular debris on the agar surface.
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5. All primary broth cultures or broth subcultures showing turbidity or pH changes should be frozen at -70°C for further testing and characterization. Agar plates showing mycoplasma colonies should be subjected to epi-immunofluorescence testing (Gardella et aL, 1983), using conjugated antisera to those mycoplasmas found frequently in cell culture infections. (See Chapters Fl and F6 in this volume.)
Minimal Solid Medium Technique This technique is considered minimal for two reasons: the DM-1 medium is probably one of the simplest used for mycoplasma isolation and only a single dish of solid medium is inoculated with the test sample. This is a departure from what is usually recommended and therefore requires some explanation. In the first large-scale study of mycoplasma contamination of cell cultures (Del Giudice and Hopps, 1978), the usual procedure of incubating all plates aerobically and anaerobically and going through an aerobic and anaerobic broth enrichment step were compared. All of the mycoplasmas that were detected on cell-free media were detected by direct inoculation and anaerobic incubation of a single agar plate. These results differ from those reported by McGarrity et al. (1979), who found that broth enrichment increased their isolation rate. The difference may be explained by differences in plating efficiency of the media used in the two studies. DM-1 medium (Gardella and Del Giudice, 1995) is a refinement of media used earlier (Del Giudice and Gardella, 1984) and was developed during attempts to grow M. hyorhinis cultivar a strains (Gardella and Del Giudice, 1995). DM-1 is a single peptone medium that is devoid of fresh yeast extract. For solid DM-1 medium, agarose is used instead of agar. Washed purified agar such as Nobel agar cannot be substituted for agarose in the DM-1 formulation.
Materials
DM'I Solid Medium Horse serum, sterile (Whittaker Bioproducts, Walkersville, MD) Distilled water SeaPlaque agarose (FMC Bioproducts, Rockland, MA) CMRL-1066 powder (Formula No. 78-5156EF, Life Technologies, Gaithersburg, MD)
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HEPES (Research Organics, Cleveland, OH) NaCl Myosate (BBL Microbiology Systems, Cockeysville, MD)
Test Strains for Quality Control Strains adapted to cell culture and stored at -70°C, as infected BHK-21 cell cultures: M. hyorhinis, BTS-7; M. orale, CH 19299; M. pimm, 70-159; M. arginini, G230; M. fermentans, PG-18; M. pulmonis, PG-34; A. laidlawii, PG-8; M. pneumoniae, FH Broth cultures stored at -70°C: M. faucium, DC333; M. lipophilum, MaBy Additional materials are the same as listed for SP-4 medium
Procedure
DM'I Medium Preparation 1. Dissolve CMRL-1066 powder (packaged for 10 liters) in 5000 ml of distilled water. This is one-half the volume of water specified on the package. Add 47.6 g HEPES and 9.35 g NaCl. 2. Adjust the pH to 7.3 and filter sterilize (220 nm). Store this 2x CMRL in the refrigerator in 500-ml amounts. 3. Dissolve 10 g of myosate and 12 g of agarose in 400 ml of distilled water. Autoclave at 12rC for 15 minutes. Cool the autoclaved solution to about 50°C, and combine with 500 ml of 2x CMRL and 100 ml of horse serum (both ingredients also warmed to 50°C). 4. Aseptically dispense medium in 5-ml amounts in petri dishes and allow to solidify. Seal the plates in plastic bags to prevent drying and store at 4°C. The DM-1 plates have a shelf life of 8 weeks.
Quality Control It is important that solid medium used for isolation has a high plating efficiency. Therefore, quality control should include titration of mycoplasma suspensions onto the solid medium to determine the highest dilution to produce colonies.
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1. Prepare test batches of DM-1 medium with each new lot of agarose, myosate, horse serum, or CMRL-1066. 2. Inoculate 10-fold dilutions of the battery of quality control strains with known titers onto medium plates and compare to DM-1 prepared with previously tested components. 3. A new lot of material is acceptable if there is no more then a 10-fold difference between the number of colonies on test and control media plates.
Mycoplasma Isolation 1. Inoculate one DM-1 medium plate with 0.1 ml of the cell culture sample, and incubate anaerobically at 36°C. 2. Microscopically examine the plate at 5 and 14 days. Most isolates develop colonies in 5 days. 3. Identify colonies by immunofluorescence (Gardella et al., 1983). Plates that show growth at 5 days may be subcultured by the push-block method for later immunofluorescence testing. 4. In order to detect cultivar a strains of M. hyorhinis, a portion of the cell culture sample should be inoculated into an indicator cell culture (Del Giudice and Hopps, 1978; Chapter F3, this volume).
Detection of Mycoplasma Contamination of Bovine Serum Although improvements in the collection and treatment of bovine serum by commercial sources might be expected to reduce the incidence of mycoplasma contamination from this source, M. arginini and other mycoplasmas of bovine origin are still prevalent contaminants of cell cultures. The chance for contamination through this means can be reduced by heat inactivation (see Chapter F5, this volume). If for some reason heat inactivation of serum is not possible, then the serum should be tested by the large volume method (Barile and Kern, 1971).
Materials A 100-ml sample of uninactivated fetal bovine serum is minimal, but five 100-ml samples yield better detection results Strains used to test for growth support: M. arginini^ G230; M. boviSy Donetta; A. laidlawii, PG8
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Fiter-sterilization (450 nm) apparatus Sterile 500-ml bottles DM-1 liquid medium, minus the horse serum component
Procedure 1. Prior to testing large volumes of bovine serum, check the sterility and ability of serum to support mycoplasma growth. 2. Using pressure sterile filtration (450 nm), aseptically dispense 400 ml of the DM-1 broth medium, without serum, into a 500-ml bottle(s). 3. Inoculate one 100-ml sample of fetal bovine serum into 400 ml of medium and incubate for 21 days at 37°C. 4. After incubation for 5 and 10 days, subculture about 0.1 ml to each of two agar plates. Incubate one plate anaerobically and one plate aerobically at each sample time. Seal the aerobic plates and examine all plates after incubation for 5 and 14 days. For quality control, a portion of the DM-1 base liquid medium is supplemented with 20% of newborn calf serum. This batch of serum must be extensively tested to ensure that it is free of mycoplasma contamination and it should be in sufficient quantity to last for an extended period of time. Challenge mycoplasma strains for the quality control test should be diluted so that less than 100 colony-forming units (CPU) are contained in the inoculum. The main limitation of the test for mollicutes in bovine serum is the low numbers of mycoplasmas usually found in contaminated serum. Mycoplasmas may be present in any particular lot of serum but may not be detected because of inadequate sample size. Thus, negative test results do not provide absolute assurance that the test serum is free of mycoplasmas.
Discussion A variety of medium formulations are suitable for use in cell culture testing provided that undefined components are performance tested before use. The formulations for the complex SP-4 and the refined DM-1 represent divergent approaches to medium development. Both media perform equally well when compared in attempts to isolate most mollicutes from cell cultures, with the exception of M. hyorhinis cultivar a strains. Kotani et al. (1990) suggested that a minor modification of conventional mycoplasma medium permitted growth of
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"noncultivable" strains of M. hyorhinis. These results cannot be reconciled with those reported by Del Giudice et al. (1980) and Gardella and Del Giudice (1995). Until some future resolution of this problem, it would be unwise to rely on axenic techniques exclusively to isolate all strains of M. hyorhinis. Detection of mycoplasmas in cell cultures is relatively easy when compared to isolating mycoplasmas from clinical materials. The gross microbial contamination of clinical materials is not present so there is no problem of microbial overgrowth obscuring the presence of a mycoplasma. Furthermore, microbial inhibitors, which may also inhibit some mycoplasmas, are not used in media for cell culture testing. Mycoplasma contaminants grow to high titer in cell cultures and when inoculated onto a mycoplasma medium usually produce abundant and easily detectable growth. Failure to detect mycoplasmas in infected cell cultures by cultural techniques may involve a number of critical factors. Probably the most frequent condition responsible is the presence of antibiotics in the cell culture sample. When cell cultures are maintained on medium containing antibiotics, it is usually recommended that they be passed two to five times on antibiotic-free medium prior to mycoplasma testing. It is difficult to appreciate the logic of using the required technique for successful aseptic passage of cells in order to provide a sample for mycoplasma test and then resume faulty techniques after the test is complete. The use of antibiotics to compensate for faulty aseptic technique is the root cause of the lateral spread of mycoplasma contamination (Chapter F6, this volume).
References Barile, M. F., and Kern, J. (1971). Isolation of Mycoplasma arginini from commercial bovine sera and its implication in contaminated cell cultures. Proc. Soc. Exp. Biol. Med. 138, 432-437. Del Giudice, R. A., and Gardella, R. S. (1984). Mycoplasma infection of cell culture: Effects, incidence, and detection. In '"Use and Standardization of Vertebrate Cell Cultures" (M. K. Patterson, ed.). In Vitro Monogr. No. 5, pp. 104-115. Tissue Culture Assoc, Gaithersburg, MD. Del Giudice, R. A., and Hopps, H. E. (1978). Microbiological methods and fluorescent microscopy for the direct demonstration of mycoplasma infection of cell cultures. In "Mycoplasma Infection of Cell Cultures" (G. J. McGarrity, D. G. Murphy, and W. W. Nichols, eds.), pp. 57-69. Plenum, New York. Del Giudice, R. A., Gardella, R. S., and Hopps, H. E. (1980). Cultivation of formerly noncultivable strains of Mycoplasma hyorhinis. Curr. Microbiol. 4, 75-80. Gardella, R. S., and Del Giudice, R. A. (1995). Growth of Mycoplasma hyorhinis cultivar a on semisynthetic medium. Appl. Environ. Microbiol. 61, 1976-1979. Gardella, R. S., Del Giudice, R. A., and Tully, J. G. (1983). Immunofluorescence. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.), Vol. 1, pp. 431-439. Academic Press, New York. Kotani, H., Butler, G. H., Tallarida, D., Cody, C , and McGarrity, G. J. (1990). Microbiological cultivation of Mycoplasma hyorhinis from cell cultures. In Vitro Cell Dev. Biol. 26, 91-96.
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McGarrity, G. J., and Barile, M. F. (1983). Use of indicator cell lines for recovery and identification of cell culture mycoplasmas. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.), Vol. 2, pp. 167-172. Academic Press, New York. McGarrity, G. J., and Kotani, H. (1985). Cell culture mycoplasmas. In "The Mycoplasmas" (S. Razin and M. F. Barile, eds.), Vol. 4, pp. 353-390. Academic Press, New York. McGarrity, G. J., Sarama, J., and Vanaman, V. (1979). Factors influencing microbiological assay of cell-culture mycoplasmas. In Vitro 15, 73-80.
F3 DETECTION OF MYCOPLASMAS BY DNA STAINING AND FLUORESCENT ANTIBODY METHODOLOGY Gerald K. Masover and Frances A. Becker
Introduction Fluorescence staining permits direct visualization of an individual mycoplasma organism. However, an individual mycoplasma is so small and so simple that even when being observed as a fluorescing body, one's confidence that what is being seen is actually a mycoplasma requires enhancement by the context in which it is being viewed. Thus, there is dependence on the nature and extent of associations between mycoplasmas and their host cells as well as on the numbers of such associations one is able to observe. The diagnostic value of a fluorescent preparation is related to the specific affinity of the stain. In the case of DNA-binding fluorochromes (DNAF), when a mycoplasma is stained it appears as a small fluorescent body because it contains DNA. The DNAF stain is not specific for mycoplasma as it will stain any body that contains DNA. Mycoplasmas also stain with specific fluoresceinated antibodies. This is sufficient by itself to diagnose mycoplasmas and to identify the isolate, if the context in which it is seen fulfills certain criteria, including: (1) a stained preparation in which the major elements are clearly resolved, (2) a sufficient number of specifically fluorescent bodies, (3) fluorescent bodies which are cell associated and arrayed in familiar patterns, and (4) presence of little or no artifactual material in the preparation. It is possible to apply both of the stains, a DNAF and a fluorescent antibody, to the same preparation and observe that a 419 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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particular small body binds both stains, thus providing diagnostic power unavailable with either stain by itself. The just-mentioned reagents can be applied directly to the cell culture sample to be tested for mycoplasma or, alternatively, to a cell culture sample which had been inoculated into a known mycoplasma-free indicator cell culture. For other information on the application of immunofluorescence techniques to detection of mycoplasmas in cell culture infections, see Del Guidice and Hopps (1978) and McGarrity et al. (1983).
Reagents Fixatives For DNAF, use 1 part glacial acetic acid plus 2 parts absolute methanol prepared fresh for each use. Prepare at least 10 ml for each coverslip to be stained. Acetone is commonly used as a fixative for immunofluorescence techniques. If acetone is used, indicator cell coverslips must be removed from their plastic dishes to a glass staining jar for fixation and then returned to the plastic dishes for the rest of the procedure. An alternative is to use 100% ethanol as the fixative. Ethanol (100%) is used for the double stain because it provides acceptable fixation without compromising the fluorescence of either of the stains (Freiberg and Masover, 1990). Stains The DNAF method will be described using bisbenzimide as the stain. The bisbenzimide concentrated stock solution (lOOOx) contains 5.0 mg bisbenzimide [Hoechst 33258, 2'-(4-hydroxyphenyl)-5-(4-methyl-l-piperazinyl)-2,5'-bi-l^benzimidazole] per 100 ml distilled water (50 jxg/ml). Each 100 ml is preserved with 0.01 g thimersal. Store this solution at 2°-8°C in the dark (foil wrapped). The bisbenzimide stain working solution (0.05 jxg/ml) contains 0.1 ml of the concentrated stock solution plus 99.9 ml citrate-phosphate buffer, pH 5.5. It is prepared fresh for each use. Fluoresceinated antimycoplasma antibodies are prepared and diluted according to well-established procedures (Gardella et al, 1983). These antibodies are typically species specific. The Evans blue counterstain, 1% stock solution (20 x) contains 1.0 g Evans blue powder (Chroma-Gesellschaft) in 100 ml phospahte-buffered saline (PBS), pH 7.2. The Evans blue working solution is 1 part of the 1% stock solution plus 19 parts PBS, pH 7.2.
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Buffers
The citrate-phosphate buffer, pH 5.5, is prepared by mixing 44 ml of 0.1 M citric acid with 56 ml of 0.2 M dibasic sodium phosphate. The phosphate-buffered saline, pH 7.2, contains 8.5 g sodium chloride, 1.1 g dibasic sodium phosphate (anhydrous) and 0.32 g sodium phosphate, monobasic, monohydrate in sufficient deionized water to make 1 liter. Mounting Solutions
The final mounting solution for DNAF stain consists of equal portions of glycerol and citrate-phosphate buffer, pH 5.5, to make the desired final volume (2 or 3 drops per slide are required). Add n-propyl gallate to make a final concentration of 0.2% (w/v). The n-propyl gallate is used to reduce photobleaching (Giloh and Sedat, 1982). This allows a particular area of a slide to be viewed for a longer period or at several different times. It also enhances the maintenance of fluorescence on storage and facilitates photography. The final mounting solution for the double stain and immunofluorescence consists of equal parts of glycerol and PBS, pH 7.2, with n-propyl gallate at a final concentration of 0.2% (w/v). The mounting solutions are stored at -10°C or below.
Procedures DNAF staining TEST SAMPLES
Testing is done on a suspension of cells in culture fluid. Passage cell cultures at least once without antibiotics or selective agents prior to testing and test the cell cultures at least 3 days after the last medium change. If an adherent cell culture is to be tested, resuspend a few cells in the culture medium by scraping them off their growth surface but do not trypsinize them. Laboratories that routinely test cell cultures for mycoplasmas may want to consider freezing samples at <-60°C as soon as they are received. The advantage of freezing samples is that groups of samples may be tested on a predetermined schedule and aliquots may be stored for retest. PREPARATION OF INDICATOR CELLS
Cells chosen as indicator cells should grow attached to a substrate and have a large ratio of cytoplasmic to nuclear area. Vero and 3T6 cells are frequently used. Cells are best seeded the day prior to inoculation of a test sample and in a
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density that will result in subconfluent growth at the end of the 3- to 5-day culture period. Prepare multiwell cell culture plates or petri dishes containing one presterilized 22-mm2 No. 1 coverslip per well (or plate). Trypsinize an antibiotic-free stock culture of Vero cells and prepare a cell suspension containing 5 x 10"^ viable cells per ml. For 6-well plates, dispense 4.0 ml of cell suspension per well (ca. 3.8-cm diameter). Adjust volume of cell suspension proportionately for different sizes of culture vessels. Incubate the coverslip-cell cultures at 35°-37°C in a humidified 4-6% CO2 air atmosphere. INOCULATION OF INDICATOR CELLS
Indicator cell cultures are inoculated 1 day after the cells are plated by adding the following to two wells each: (a) Test sample, 0.5 ml. Multiple samples may be tested concurrently, (b) One hundred colony-forming units (CFU) or less of positive control Mycoplasma hominis (derived from ATCC No. 23114) in 0.5 ml or less of complete mycoplasma broth. The intent of this positive control is to test a low number of poorly cytadsorbing mycoplasmas. Therefore, M. ovale strains or other poorly cytadsorbing mycoplasma species may also be used but they may not give reliably positive results when inoculated in low numbers, (c) One hundred CFU or less of positive control M. hyorhinis, strain BTS-7 (derived from ATCC No. 17981), in 0.5 ml or less of complete mycoplasma broth. The intent of this positive control is to test a low number of strongly cytadsorbing mycoplasmas. Therefore, other strains of M. hyorhinis or other cytadsorbing mycoplasmas may be used, (d) Negative control, uninoculated wells. DNAF STAINING
1. Three to 5 days after test sample addition, remove medium from the well (do not allow the coverslip to dry). 2. Add fixative in sufficient volume to completely immerse the coverslip (2 to 5 ml) and let stand for 2 minutes. 3. Aspirate the fixative and then apply fresh fixative. Let stand for 5 minutes. 4. Aspirate the fixative and air dry (30-60 minutes) or aspirate fixative and rinse twice with citrate-phosphate buffer, aspirating after each rinse. 5. Apply working solution of bisbenzimide stain (sufficient quantity to immerse the coverslip) and stain for 10-30 minutes in the dark. 6. Aspirate the stain and wash two times with distilled water. 7. Aspirate the distilled water and either mount or store if desired. Stained coverslips may be stored refrigerated for up to 7 days either dry or immersed in citrate-phosphate buffer. 8. Mount the coverslips on a slide with cell side down, on a drop of mounting solution. A more permanent mount may be made by sealing the mounted coverslip with petroleum jelly, nail polish, or an equivalent liquid barrier. Coverslips
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mounted in this manner and stored refrigerated in the dark may be viewed within several weeks. 9. Observe microscopically (minimum 630X; oil immersion). 10. Microscopy: Use a microscope equipped with a fluorescence light source, an epi-illumination system, and fluorite objectives. Filters with characteristics similar to those given are suitable for bisbenzimide: Excitation Barrier Beam splitter
BP365/11 (bandpass 365 nm; U-nm bandwidth) LP397 (long-wave pass 397 nm) FT395 (dichromatic beam filter 395 nm)
INTERPRETATION OF RESULTS
The bisbenzimide-DNA complex emits 475 nm wavelength light, which is a light blue color. The indicator cell nuclei are intensely stained with entire margins and are generally ovoid. Mitochondrial DNA is not readily visualized with this stain and, therefore, cell cytoplasmic areas are not observed. The background on a good preparation is black. Occasionally, micronuclei formed by anomalous mitotic events are seen but they are usually much larger than mycoplasmas. Occasional mitotic figures will also be observed. The nuclei are approximately 10 to 20 \xm in diameter (or long axis) and the mycoplasmas are seen as small round bodies approximately 0.3 iJim in diameter. The mycoplasmas are generally not intracellular but are adherent to the cell membrane and are seen in the areas where the cytoplasm is expected to be. When the indicator cells are well separated, heavily contaminated cells have a blue fluorescent nucleus with the cytoplasmic region virtually covered with discrete, small, blue fluorescent bodies and the space between the cells will generally be black. Mycoplasmas also cover the membrane above the nucleus but these are not distinguishable from the very brightly fluorescing nucleus. The extent of cytadsorption is variable with different mycoplasma species and in some preparations, small blue fluorescent bodies are seen adhering to the glass slide between cells. These cannot be confidently diagnosed as mycoplasmas by use of bisbenzimide alone (refer to the double stain procedure in this chapter). The major features of DNAF staining are illustrated in Fig. la. For a test to be considered valid, both negative control slides must be negative for mycoplasma and at least one positive control slide for each of the positive control mycoplasma species must be positive. We have established in our laboratory that for a test sample to be considered positive for mycoplasma, at least 5% of not less than 200 cells in a valid test must have multiple (more than 5) blue fluorescent bodies the size of mycoplasmas associated with the indicator cells. In the event of an ambiguous result or a determinate error, a retest should be performed. Some samples, notably hybridoma cultures, contain particulate debris that reacts with the stain rendering
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the test result ambiguous. In such cases a retest is done in which 0.5 ml of supernatant medium and indicator cells resuspended by scraping from the original culture is subcultured into fresh indicator cultures after 3 to 5 days of incubation. The subculture is stained and observed after an additional 3 to 5 days of incubation. Immunofluorescent Antibody Staining
Preparation of the indicator cells, specimen handling, and inoculation of the indicator cells are the same for this method as for the DNAF method. Each test should include both negative and positive controls. Ideally, positive controls corresponding to each antibody preparation used in a cocktail should be included in each test. Alternatively, antibody-positive-control testing should be done at some reasonable interval to assure that each antibody preparation in the cocktail is working. 1. Three to 5 days after inoculation of the indicator cell culture with sample, aspirate medium and fix for 5 minutes (100% ethanol or acetone). Do not allow cell preparations to dry. Aspirate the fixative and replace with fresh fixative for an additional 20 minutes, then rinse with two changes of PBS, pH 7.2. 2. Stain by adding sufficient diluted fluoresceinated antibody to cover the indicator cell culture (approximately 0.5 ml) and allow to stain for 30 minutes. Rinse twice with PBS and aspirate fluids. 3. Counterstain with Evans blue working solution for 30 minutes. Rinse twice with PBS and aspirate fluids. 4. Mount the coverslips cell side down as described for the DNAF method but using the PBS/glycerol mounting solution. 5. Observe microscopically (minimum 630X; oil immersion). 6. Microscopy: Use a microscope equipped with a fluorescence light source, an epi-illumination system, and fluorite objectives. Filters with characteristics similar to those given below are suitable for fluorescein: Excitation Barrier Beam splitter
BP485/20 (bandpass 485 nm; 20-nm bandwidth) LP 520 (passes light above 520-nm wavelength) FT 510 (dichromatic beam filter 510 nm)
INTERPRETATION OF RESULTS
Specific fluorescein fluorescence is apple green, whereas nonspecific fluorescence is generally yellow. The Evans blue counterstain gives the entire cell a deep red color which provides a striking background for the mycoplasmas and eliminates cell-associated nonspecific fluorescence. The mycoplasmas are seen as small (ca. 0.3-|jLm diameter) green round bodies, generally cell associated. They are seen singly and in groups on both the cytoplasmic and nuclear areas of
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the cell. Observation of specific fluorescence with fluoresceinated antibody preparations used singly is sufficient to identify mycoplasmas at the species level. A minimum number of green fluorescent bodies on a minimum number of cells may be defined as a diagnostic minimum requirement as described for DNAF earlier. In the case of mycoplasmas that cytadsorb well, there is rarely question about whether or not there is an infection. However, for those mycoplasmas that cytadsorb poorly, it is best to systematically observe a minimum of about 200 cells. The major features of immunofluorescence staining are illustrated in Fig. lb. The apple green color of specific fluorescence is easily compromised in the photographic process but the characteristic green color should be seen using the microscope. IMMUNOFLUORESCENCE STAINING OF AGAR CULTURES
The principle use of this method is serological identification of mycoplasmas isolated on agar. The technique involves application of the fluoresceinated antibodies directly to the agar culture and visualization of fluorescing colonies by fluorescence microscopy. Gardella et al. (1983) and Barile et al. (1973) describe the method in detail.
Double Stain Method: Combined DNAF and Immunofluorescence
Preparation of the indicator cells, specimen handling, and inoculation of the indicator cells are the same for the double stain method as for the DNAF alone. Each test should also include both positive and negative controls. The staining method is illustrated diagrammatically in Fig. 2. 1. Three to 5 days after the addition of test samples, remove the supernatant cell culture medium and rinse the cell layer twice with PBS. 2. Fix by rinsing the coverslip cell culture twice with 100% ethanol and then adding fresh 100% ethanol for an additional 20 minutes. 3. Remove the ethanol by aspiration and rinse twice with PBS, pH 7.2. 4. Add about 0.5 ml of the working dilution of fluoresceinated antimycoplasma antibody preparation and incubate at room temperature for 30 minutes. One or more antibody preparations against different mycoplasma species may be combined in this procedure. We have used mixtures of up to five different antisera. 5. Remove the antibodies by aspiration and rinse twice with PBS, pH 7.2. 6. Add the bisbenzimide working solution in sufficient quantity to immerse the coverslip and stain in the dark for 5 minutes. Remove the bisbenzimide by aspiration and add fresh bisbenzimide working solution. Allow to stain in the dark for an additional 15 minutes.
Fig. 2.
426
Double stain procedure.
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7. Remove the bisbenzimide stain by aspiration and rinse twice with PBS, pH 7.2. 8. Add the Evans blue counterstain and allow to stain at room temperature for 30 minutes. Remove the Evans blue by aspiration and rinse twice with PBS. 9. Mount the coverslips cell side down as described for the DNAF method but using the PBS/glycerol mounting solution. 10. Observe microscopically (minimum 630x; oil immersion). For the double stain, a field of interest is selected using one of the filter sets (bisbenzimide or fluorescein; see later) and then the same field is observed using the other filter set. This is done by sliding the filter housing to the second position without making any changes in the field of observation. It is useful to continuously make minor adjustments of focus and to switch the filter sets back and forth several times for each field being observed. 11. Microscopy: Use a microscope equipped with a fluorescence light source, an epi-illumination system, and fluorite objectives. For the double stain method, the microscope must also be equipped with a carriage for fluorescence filter sets which is capable of holding at least two filter sets. Suitable filter sets for both bisbenzimide and fluorescein have been described previously. INTERPRETATION OF RESULTS
The bisbenzimide and fluorescein stains give results as described earlier. Occasionally, in some preparations, it is possible to faintly see the red counter stain (Evans blue) used to enhance the fluorescein stain while using the DNAF filter set. This does not interfere with evaluation of the DNAF. The major advantage of this method is the ability to switch back and forth between filter sets so that a particular small cell-associated body may be seen to stain both blue with the DNAF stain (contains DNA) and green with the fluorescent antibody stain (contains specific mycoplasma antigens). This information, in addition to the size, shape, and cell association, serves to greatly enhance confidence in the specific diagnosis of mycoplasma. On some preparations, small mycoplasmasized bodies are seen adherent to the glass slide away from host cells. The double stain makes diagnosis of these as mycoplasmas much more certain. It is necessary to focus on a particular body or small array of individual bodies and satisfy yourself that they stain with both stains and are the right size and morphology in order to define them as mycoplasmas. By use of individual fluorescent antibody preparations, identification of the mycoplasma can be made. By use of several fluoresceinated antibodies in the same preparation, i.e., against the five most common cell culture contaminants, it is possible to rapidly screen the cell cultures for mycoplasma contamination. A cocktail of fluoresceinated antibody preparations against M. hyorhinis, M. orale, M. fermentans, M. arginini, and Acholeplasma laidlawii would be expected to
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detect 95% of the cell culture contaminant mycoplasmas (Barile and Rottem, 1993). It is also possible to apply the double stain directly on the test culture (without indicator cells) and have a reasonably confident result in about 3 hours even in the presence of various types of debris and artifacts. The major features of the double stain can be seen by comparing Figs, la and lb, showing the same field photographed with the DNAF and immunofluorescence filter sets, respectively. The figure shows that the putative mycoplasmas that stain blue in the DNAF stain (Fig. la) match up with green fluorescent bodies in the fluorescent antibody stain (Fig. lb), thus demonstrating that the bodies contain both DNA and mycoplasma antigens and are of approximately the size and shape of a mycoplasma.
Discussion Although fluorescence microscopy permits visualization of a single mycoplasma cell, it will be necessary to view a larger number of mycoplasma cells in order to achieve reliable diagnosis. For example, we have established five fluorescent bodies in 5% of not less than 200 cells as a minimum for a positive test using DNAF. Most often cell culture contaminant mycoplasmas are present in high concentrations and produce infections that are readily detectable by use of these methods. The reliability of the methods depends on control of the procedure, reagents, and equipment and careful observation of the stained preparations by a well-trained observer. The methods described in this chapter involve the use of indicator cells. However, cell cultures can be tested directly without the use of indicator cells. In this case, cells are prepared for staining by growing them on coverslips, scraping cells from the monolayer culture, and drying them onto a slide or centrifuging cells onto a slide with, for example, a Cytospin centrifuge. The direct method has the advantage of speed and simplicity: a direct DNAF, fluorescent antibody, or double stain preparation can be completed and ready to view in less than 3 hours. In some of the direct DNAF preparations, however, artifacts originating from extracellular DNA of damaged or dead cells reduce the confidence of diagnosis. The absence of normal and consistent cell morphology against which to view contaminant mycoplasmas is also a disadvantage of direct tests using either DNAF or fluorescent antibody. The use of both stains together in such cases assists greatly in distinguishing between mycoplasmas and artifacts. The use of indicator cells improves the reliability of the system substantially by permitting standardization of the culture system and allowing for inclusion of positive and negative controls. It also increases the sensitivity of the test by providing a biological system capable of detecting mycoplasmas that grow poor-
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ly, or not at all, in conventional broth and agar media (see Chapter F2, this volume). In theory, a single viable mycoplasma can be amplified and the progeny visualized; in practice this is probably achieved only with strongly cytadsorbing species such as M. hyorhinis. With the use of indicator cells, the fluorescent methods are relatively rapid, requiring only 3 to 5 days to complete, a fraction of which is hands-on time. Although some Mycoplasma species cytadsorb poorly, there are usually sufficient numbers to provide a definitive diagnosis. However, for this reason, mycoplasma screening programs should always also include standard cultural methods.
References Barile, M. F., and Rottem, S. (1993). Mycoplasmas in cell culture. In "Rapid Diagnosis of Mycoplasmas" (I. Kahane and A. Adoni, eds.), pp. 155-193. Plenum, New York. Barile, M. F., Hopps, H. E., Grabowski, M. W., Riggs, D. B., and Del Giudice, R. A. (1973. The identification and sources of mycoplasmas isolated from contaminated cell cultures. Ann. N. Y. Acad. Sci. 225, 251-264. Del Giudice, R. A., and Hopps, H. E. (1978). Microbiological methods and fluorescent microscopy for the direct demonstration of mycoplasma infection of cell cultures. In "Mycoplasma Infection in Cell Cultures" (G. J. McGarrity, D. G. Murphy, and W. W. Nichols, eds.), pp. 57-69. Plenum, New York. Freiberg, E. F., and Masover, G. K. (1990). Mycoplasma detection in cell culture by concomitant use of bisbenzimide and fluoresceinated antibody. In Vitro Cell Dev. Biol. 26, 585-588. Gardella, R. S., Del Giudice, R. A., and TuUy, J. G. (1983). Immunoflurorescence. In "Methods in Mycoplasmology" (S. Razin and J. G. Tully, eds.). Vol. 1, pp. 431-439. Academic Press, New York. Giloh, H., and Sedat, J. W. (1982). Fluorescence microscopy: Reduced photobleaching of rhodamine and fluorescein protein conjugate by n-propyl gallate. Science 217, 1252-1255. McGarrity, G. J., Steiner, T., and Vanaman, V. (1983). Detection of mycoplasmal infection in cell cultures by DNA fluorochrome staining. In "Methods in Mycoplasmology" (J. G. Tully and S. Razin, eds.). Vol. 2, pp. 183-190. Academic Press, New York.
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F4 DETECTION OF MYCOPLASMA INFECTION BY PCR Connie Veilleux^ Shmuel Razin^ and Laurie H. May
Introduction The polymerase chain reaction (PCR) is a rapid and sensitive assay that can be used routinely to screen for mycoplasmas in cell culture samples (Teyssou et al., 1993; Wong-Lee and Lovett, 1993; Spaepen et al., 1992). The assay described here uses universal primers that amplify a conserved region of the 16S ribosomal RNA gene found in all prokaryotes (Weisburg et al., 1989). The use of these primers in a PCR assay allows the detection of any bacterial infection in cell cultures. Because of the high degree of amplification achieved by PCR, stringent quality control practices such as those described by Kwok and Higuchi (1989) are essential throughout sample preparation and amplification in order to avoid false-positive results.
Materials Note: It is essential that the first seven items listed below be free of bacterial DNA. These reagents should be tested to ensure the absence of bacterial DNA. Testing may be done by either the vendor or the user against a panel of positive and negative controls. The heating block of the PCR machine needs to be cleaned periodically with a 10% (v/v) solution of bleach. 431 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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Distilled and deionized water (Mallinckrodt H453) or irrigation grade: Use in preparation of sample buffer, proteinase K, phosphate-buffered saline (PBS), master mix solutions, and primers. This high purity water is not needed for the electrophoretic analysis of the PCR products (items 18-24 on this list). Prokaryotic 16S rDNA primers: A 780-bp product Nucleotide position
Nucleotide sequence, 5' to 3'
Length (bases)
21 802
GAT CCT GGC TCA GGA TGA AC GGA CTA CCA GGG TAT CTA ATC
20-MER 21-MER
Sample buffer: 50 mM KCl, 10 mM Tris-HCl, pH 8.3, 0.45% (v/v) Nonidet P-40 (NP-40), and 0.45% (v/v) Tween 20 PBS, pH 7.2 0.2 Proteinase K (molecular biology grade): stock solution 10 mg/ml Stock solutions for master mix (store at -20°C): 100 mM Tris-HCl, pH 8.3, 500 mM KCl 10 mM deoxynucleotide triphosphates (dNTPs) 50 |xM primers 15 mM MgCl2 (store at 4°C): Vortex vigorously for 2 minutes before use 5 U/|xl AmpliTaq DNA polymerase: highest purity available (Perkin-Elmer Norwalk, CT), vortex before use Mineral oil (molecular biology grade) Sterile 2.0-ml microtubes with attached screw caps (version C: Sarstedt, Newton, NC) for sample preparation. Screw-cap tubes reduce the formation of aerosols. Sterile 0.5-ml microcentrifuge tubes for amplification DNA Thermal Cycler 480 (Perkin-Elmer): The profile used is an initial denaturation step at 94°C for 2 minutes; 24 cycles at 94°C for 1 minute, 55°C for 1 minute, 72°C for 1 minute; and an extension step at 72°C for 7 minutes. The final step is at 4°C until tubes are removed from block. Note: Every thermal cycler is different. Modifications must be made to the times and cycle number if another thermal cycler is used. Assay negative control: A known uncontaminated cell line containing less than 1 X 10^ cells/ml in culture medium. Store at -20°C in 1-ml portions. Mycoplasma DNA used for sample positive control and assay positive control: A suspension containing 10^ colony-forming units (CPU) per ml of washed Acholeplasma laidlawii (ATCC No. 29804) cells in 10 mM Tris-HCl (pH 7.5) and 0.1 mM EDTA. Store at -20°C in 75- to 110-|xl portions. Assay positive control: A cell line diluted to less than 1 x 10^ cells/ml in culture medium. Mycoplasma DNA is diluted 1:10 into the cell suspension. Place 1-ml portions into screw-cap tubes. Store at -20°C.
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Microcentrifuge (14,000 rpm/min, equivalent to 9000 g) Water bath at 55°C Vortex Dry bath at 95°C 95% and 100% ethanol lOx TBE buffer (1 liter): 108 g Tris-base, 55 g boric acid, 40 ml 0.5 M EDTA, pH8.0 3% NuSieve GTG agarose gel: Add 2 g NuSieve (FMC Bioproduct, Rockland, ME) and 1 g agarose to 100 ml of 1 x TBE buffer. Use either a boiling water bath or a microwave oven to liquify the agarose. Cool to 65°C before pouring gel. After gel has solidified for 1 hour at room temperature, store at 4°C in 1 x TBE buffer until samples are loaded. Salt buffer: 100 mM Tris-HCl, pH 7.5, 100 mM MgCl2, 1 M NaCl, 10 mM dithiothreitol Loading dye solution: 0.25% bromophenol blue in 40% sucrose (w/v) DNA molecular weight marker VIII (Boehringer Mannheim) Ethidium bromide: stock solution (10 mg/ml) Shaker Ultraviolet (UV) transilluminator for analytical gels Photographic apparatus
Procedure
Cell Culture Sample Preparation
1. Dilute proteinase K to a final concentration of 60fxg/jjLl in sample buffer. 2. Thaw one assay positive control and one assay negative control at room temperature (see Table I). 3. Three microtubes are required for each cell culture sample tested: Add 1 ml of sample suspension per tube. 4. Centrifuge samples and assay controls at maximum speed (9000 g) in a microcentrifuge for 30 minutes 5. Decant supernatant, add 500 JJLI of PBS, and vortex. Centrifuge at maximum speed in a microcentrifuge for 15 minutes. Complete a total of two PBS washes and three centrifugations. Always prepare the assay positive control last and change gloves before the next step is performed. It is not necessary to remove all the PBS at each wash. 6. Decant the PBS. Add 450 jxl of sample buffer with the proteinase K and vortex. [Optional: If there is a large pellet make a 1:10 dilution of one sample and the sample to be used as the sample positive control. Use PBS or deionized H2O as
TABLE I PCR ASSAYCONTROLS Name
Number of control tubes
Description
Expected results
Assay negative control
One per test series
Uninfected cell line
Negative
Assay positive control
One per test series
Positive
Sample positive control
One per sample tested
Cell line infected with A . laidlawii Test sample infected with A. laidlawii
Reagent negative control
One per test series
Master mix sample with no DNA template
Negative
Positive
Purpose of control Determines whether exogenous DNA contamination occurred during sample preparation Indicates that proper procedure was followed throughout assay Determines whether inhibitors from media are present in sample preparation. Indicates whether cell population is too dense ( > l X lo5 cellslml) Indicates whether master mix reagents are free of exogenous DNA
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diluent. Set aside one tube per cell culture sample for the sample positive controls (see Table I).] 7. Incubate the remaining samples at 55°C for 1 hour or overnight. 8. To the sample positive control tubes set aside in step 6, add 50 |xl mycoplasma DNA solution and vortex. Incubate at 55°C for 1 hour or overnight. It is important to complete this step after all other tubes are prepared and closed to prevent mycoplasma DNA carryover to another sample. 9. Incubate all samples at 95°C for 10 minutes to inactivate proteinase K. Store samples at -20°C until ready for PCR amplification. PCR Amplification and Product Analysis
1. Prepare the master mix on ice. The final concentrations in the master mix are as follows: 10 mM Tris-HCl, pH 8.3, 50 mM KCl, 200 |xM dNTP, 0.5 |xM of each primer, 1.5 mM MgCl2, and 0.025 U/jjil AmpliTaq DNA polymerase. 2. Dispense 45 jxl of the master mix per 0.5-ml microcentrifuge tube. As a reagent negative control (see Table I), use 45 jxl of the master mix and 5 |xl of deionized water. Place all tubes on ice. 3. Overlay each tube with 30 |xl of mineral oil. 4. Thaw samples and briefly centrifuge to remove condensation from the cap. This needs to be performed in an area away from the master mix. Transfer 5 |xl of each sample to 45 |JL1 of the master mix. 5. Start the thermal cycler and place the tubes in the block when the temperature reads 70°-90°C. Run the thermal cycler profile described under Materials. 6. Remove samples from the thermal cycler and carefully pipette off the mineral oil. Add 2.5 vol of chilled 100% ethanol and vortex. Leave for 30 minutes at -70°C. 7. Centrifuge at maximum speed in a microcentrifuge for 30 minutes. Decant the supernatant and wash pellet with 95% ethanol. Invert tubes and allow to dry for 1 hour. 8. To each tube add 11 jxl deionized water, 1.5 |JL1 of salt buffer, and 2.5 |xl of loading dye solution. Mix well and warm to 65°C for a few minutes. 9. Prepare a solution of DNA molecular weight marker according to manufacturer's instructions. 10. Load 15 |xl of the DNA molecular weight marker and the samples in individual lanes on the gel. Run the gel at 100 V for 80-100 minutes. 11. Transfer the gel to an ethidium bromide solution (0.5 |xg/ml) and place on a shaker for 45-60 minutes. 12. Observe gel on the UV transilluminator and photograph. Sensitivity Testing
The sensitivity of the PCR method relative to the culture method needs to be established before samples are run. To do this, serial 10-fold dilutions of the
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A. laidlawii cell suspension are prepared in assay negative control samples and tested as described earlier. In our laboratory, the sensitivity of the PCR assay with this protocol is 100-1000 CFU/ml of A. laidlawii. This should be adequate to diagnose cell cultures infected with mycoplasmas since infections usually result in mycoplasma titers of 10^-10^ CFU/ml (McGarrity, 1982). The number of cycles can be adjusted to achieve the desired sensitivity. Results
The controls need to be checked first to verify that the assay worked as expected (Table I, Expected Results). The sample positive control and the assay positive control should exhibit a specific band at 780 bp as shown in Fig. 1. A cell culture sample is considered positive when a band can be seen at 780 bp (Fig. 1). If unusual results are observed, such as a band at 50 bp (primer-dimer) or a band at 780 bp in only one of the duplicates, that sample will need to be retested. A 1:10 dilution of the sample may yield better results if a component in the sample is inhibitory to the PCR. The sample positive control is an important control. A constant amount of A. laidlawii (10,000 CFU/ml) is added to each sample, which should result in a 1 2 3 4 5 6 7 8 9 10 111213141516171819 20
780 bp-
20 bp.i 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 Fig. 1. Amplification from a set of cell culture samples. (Lanes 1 and 21) Hpall and Oral plus H/ndlH DNA size markers; (lanes 2 to 5) sensitivity curve, 10, 100, 1000, and 10,000 CFU/ml of/A. /a/d/aw//; (lanes 6, 7, 9, 10, 12, 13, 15, 16, 18, 19, 22, 23, 25, 26, 28, 29, 31, 32) cell culture samples in duplicate; (lanes 8, 11, 14, 17, 20, 24, 27, 30, 33, 35, 37) sample positive controls; (lane 34) a knov^n negative cell culture sample; (lane 36) a knov^n positive cell culture sample; (lane 38) assay negative control; (lane 39) assay positive control; and (lane 40) reagent negative control.
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band of similar intensity to that seen in the sensitivity curve at the same concentration (Fig. 1). An example of an unexpected result is seen in Lane 17, in which the sample positive control appears as a weak band, similar to the 100 CFU/ml concentration in the sensitivity curve. A pipetting error or inhibition of the reaction by something in the sample may explain this result.
Discussion PCR is becoming a powerful tool in the diagnostic laboratory. The method described in this chapter allows for 10-15 samples to be prepared and assayed in 48 hours with consistent results. An advantage of the PCR assay is that it is unnecessary to purify the DNA from the samples for a screening assay of mycoplasmas in a cell culture. The sample preparation described in this chapter was designed to minimize sample handling, thereby reducing the risk of contamination from exogenous DNA sources. In addition, the sample buffers and solutions were selected because they contain no ingredients known to inhibit the PCR reaction. The series of washes with PBS were performed (Higuchi, 1989) to remove the growth medium, and the high-speed centrifugation was done to remove some cell debris as well as lyse the cells. Samples prepared in this manner can be used in assays designed to detect mycoplasmas as well as other bacteria. However, the weaknesses of the PCR technique are often overlooked and are therefore an important part of this discussion. Amplification via PCR allows for a low copy number of DNA to be detected. For this reason, care must be taken to prevent exogenous DNA from entering a sample during preparation and amplification. Therefore, it is important to adhere to strict quality control procedures to prevent false-positive results. There is value in autoclaving solutions when possible (Kwok and Higuchi, 1989) and in assuring that solutions are (1) sterile, (2) handled with an awareness of aseptic technique, and (3) expiration dated to assure that they are used within reasonable time periods. The Taq polymerase has been shown to be contaminated with 100-1000 genome equivalents of bacterial DNA per unit of enzyme during its production and purification (Rand and Houck, 1990). It is important to buy Taq polymerase from a source that removes as much of the exogenous bacterial DNA as possible. Restricting the thermal cycler profile to 25-30 cycles and the sensitivity to 100-1000 CFU/ml will also assist in reducing the appearance of false positives due to the Taq polymerase. Use of the PCR carryover prevention kit (Perkin-Elmer) or the isopsoralen component 10 (HRI, Concord, CA) is highly recommended with this assay. The introduction of an amplified product to sample is a well-known source of exogenous DNA that can be prevented by either of these methods.
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It is important to note that PCR will detect DNA of both viable and nonviable mycoplasmas. This can cause results that may be difficult to interpret when testing a cell line that has been treated with antimycoplasma drugs because such cultures may contain residual DNA from nonviable mycoplasmas. In these instances, direct cultivation or an immunoassay is recommended in addition to PCR. The use of primers that amplify the 16S ribosomal RNA gene offers the advantage of detecting any prokaryote, including mycoplasma, that infects cell cultures. For most purposes, finding that a cell culture is infected with a prokaryote is all the investigator needs to know. If specific identification of the infecting mycoplasma is desired, the 780-bp PCR product can be further digested using restriction enzymes. This restriction pattern can then be compared to those of the mollicute species known to infect cell cultures (Deng et aL, 1992).
References Deng, S., Hiruki, C , Robertson, J. A., and Stemke, G. W. (1992). Detection by PCR and differentiation by restriction fragment length polymorphism of Acholeplasma, Spiroplasma, Mycoplasma, and Ureaplasma, based upon 16S rRNA genes. PCR Methods Appl. 1, 202-204. Higuchi, R. (1989). Simple and rapid preparation of samples for PCR. In "PCR Technology: Principles and Applications for DNA Amplification" (H. A. Erlich, ed.), pp. 31-38. Stockton Press, New York. Kwok, S., and Higuchi, R. (1989). Avoiding false positives with PCR. Nature (London) 339, 237238. McGarrity, G. J. (1982). Detection of mycoplasma infection of cell cultures. In "Advances in Cell Cultures" (K. Maramorosch, ed.). Vol. 2, pp. 99-131. Academic Press, New York. Rand, K. H., and Houck, H. (1990). Taq polymerase contains bacterial DNA of unknown origin. Mol Cell. Probes 4, 445-450. Spaepen, M., Angulo, A. F., Marynen, P., and Cassiman, J.-J. (1992). Detection of bacterial and mycoplasma contamination in cell cultures by polymerase chain reaction. FEMS Microbiol. Lett. 99, 89-94. Teyssou, R., Poutiers, F., Saillard, C , Grau, O., Laigret, F., Bove, J.-M., and Bebear, C. (1993). Detection of mollicute contamination in cell cultures by 16S rDNA amplification. Mol. Cell. Probes 7, 209-216. Weisburg, W. G., Tully, J. G., Rose, D. L., Petzel, J. P., Oyaizu, H., Yang, D., Mendelco, L., Sechrest, J., Lawrence, T. G., Van Etten, J., Maniloff, J., and Woese, C. R. (1989). A phylogenetic analysis of the mycoplasmas: Basis for their classification. J. Bacteriol. 171, 6455-6467. Wong-Lee, J. G., and Lovett, M. (1993). Rapid and sensitive PCR method for identification of Mycoplasma species in tissue culture. In "Diagnostic Molecular Microbiology: Principles and Applications" (D. H. Persing, T. F. Smith, F. C. Tenover, and T. J. White, eds.), pp. 257260. Am. Soc. Microbiol., Washington, DC.
F5 ANTIBIOTIC TREATMENT OF MYCOPLASMA-INFECTED CELL CULTURES Richard A. Del Giudice and Roberta S. Gardella
Introduction Mycoplasma-infected cell cultures are best discarded and replaced with clean cultures from frozen stocks or cell culture repositories. However, if a particular cell culture is irreplaceable it should be cured of its mycoplasma infection. The treatment protocol plus monitoring, although laborious, is less time-consuming than establishing a new cell line with the required characteristics. A variety of novel techniques to cure mycoplasma-infected cell cultures have been used with variable success. Cells have been treated by passage in nude mice; have been exposed to 5-bromouracil, bisbenzimide H33258 fluorochrome and light; have been treated with serum or antiserum; and have been exposed to heat or detergents (Marcus et aL, 1980; Van Diggelen et aL, 1911 \ Barile, 1979). Problems with efficacy or toxicity are not uncommon with most of these techniques. Antibiotic treatment is the most conventional and probably the most common method but this too has met with variable success. Some procedures in the literature involve the use of antibiotics long available to the cell culturists, whereas others describe the use of a new series of antibiotics either recently evaluated for or specifically designed for the treatment of mycoplasma-infected cell cultures. Some of these newer additions include ciprofloxacin (Mowles, 439 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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1988), mycoplasma removal agent (MRA), and enrofloxacin (Fleckenstein et al., 1994). Drexler et al. (1994) reported a failure rate of 25% using antimycoplasma drugs. The problem of low efficacy was accompanied by high toxicity; 13% of the cell cultures did not survive the treatment. The reason that these poor results are typical is that susceptibility tests on the mycoplasma contaminant are not done; drugs are used in a hit or miss fashion. To some nonmycoplasmologist who might find the performance of susceptibility tests demanding, the allure of a "shotgun" cure may be tempting and in some cases worth a try. We developed this protocol to provide a service to cell culture laboratories (Gardella and Del Giudice, 1984). Using this protocol, we have cured approximately 40 cell lines, some of which were infected with two different Mycoplasma species. So far we have not received an infected cell culture that resisted treatment and no ill effects due to drug toxicity were observed. Key factors to success are (i) select drugs for minimal toxicity, (ii) perform drug susceptibility tests on the mycoplasmas isolated from the infected cell cultures, and (iii) enhance the effect of the drugs by appropriate manipulation of the cultures (Gardella and Del Giudice, 1984; Del Giudice and Gardella, 1984).
Materials Antibiotics. The panel of drugs should be periodically evaluated for performance. Ineffective drugs are replaced with drugs which show promise as antimycoplasma agents. Antibiotic stock solutions are prepared in water, filter sterilized, frozen, and stored in small volumes as follows: linocomycin hydrochloride, 4 mg/ml (Sigma); kasugamycin hemisulfate, 4 mg/ml (Sigma); neomycin sulfate, 4 mg/ml (Sigma); kanamycin sulfate, 4 and 8 mg/ml (Sigma); tetracycline hydrochloride, 4 and 2 mg/ml (Calbiochem and Sigma); tylosin tartrate, 5 mg/ml (Flow Laboratories and Sigma); ciprofloxacin hydrochloride, 4 mg/ml (Sigma); spectinomycin dihydrochloride, 4 mg/ml (Sigma); spiramycin, 4 mg/ml (Sigma) Sterile paper discs (BBL Microbiology Systems) Suitable mycoplasma agar medium in 60-mm dishes Liquid mycoplasma medium for stock cultures M-CMRL medium for M. hyorhinis cultivar a (Gardella and Del Giudice, 1995) Sterile phosphate-buffered saline, pH 7.2, in 1.8-ml quantities BBL GasPack anaerobic system (Becton-Dickinson Microbiology Systems) UV microscope with incident illumination for immunofluorescence of mycoplasma colonies growing on agar (Del Giudice et al, 1976) Fluorescent antibody conjugates directed against those mycoplasmas known to contaminate cell cultures
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Antiserum directed against those mycoplasmas known to contaminate cell cultures
Procedures
Mycoplasma Cultivation 1. Mycoplasmas are isolated from infected cell cultures by agar cultivation and are identified by immunofluorescence (Del Giudice et al., 1976). 2. A portion of the original infected culture is also inoculated into liquid mycoplasma medium to be used as a stock culture and is frozen at -70°C. For some M. hyorhinis cultivar a strains, initial plating on agar and growth in broth are accomplished in M-CMRL medium. 3. In the case of samples infected with strains of more than one mycoplasma species, portions are inoculated into tubes of liquid mycoplasma medium, each with an appropriate antiserum to allow for selective growth of one strain. 4. Seed stocks are serially diluted 10-fold and are plated onto solid medium, to enumerate colony-forming units (CFU). 5. For cultures grown in the presence of antiserum, titers on agar are also determined, and colonies obtained from final dilution plates are used to initiate stock cultures. Immunofluorescence of agar colonies is used to monitor culture purity. Antibiotic Susceptibility Testing 1. One-tenth-milliliter quantities of stock culture dilutions containing 10^ CFU/0.1 ml are inoculated onto mycoplasma agar plates. Plates are rotated to spread the inoculum evenly over the agar surface. Some strains of M. hyorhinis cultivar a are plated on M-CMRL agar (Gardella and Del Giudice, 1995). Plates are dried for 30-60 minutes at room temperature. 2. Paper disks are saturated by touching them to the surface of antibiotic solutions and are placed in the center of the inoculated plates. 3. Nine or 10 antibiotics, each at two dilutions, are used. Solutions used to wet disks consist of undiluted and a 1:10 dilution of the antibiotic solution. 4. Control plates with no drugs are similarly prepared to assess uninhibited growth. 5. Plates are incubated at 35°C aerobically or anaerobically (BBL Gaspack), depending on the atmospheric requirements of the organism.
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6. Zones of inhibition are measured when control plates show adequate growth, generally after 5 and 10 days. 7. Sixty-millimeter agar plates are divided into three equidistant concentric sectors surrounding the disks; each sector is approximately 7.0 mm. Growth in these sectors is used to estimate zones of inhibition as follows: 4+ inhibition, no growth on the plate; 3+ inhibition, growth in the outermost sector only; 2+ inhibition, growth in two outer sectors; 1 + inhibition, growth in all three sectors; and no inhibition, unaffected growth. Breakthrough is also recorded, that is when some plates show colonies in a particular sector, but growth is inferior to growth on control plates. 8. Two antibiotics are generally selected to minimize the chances of outgrowth of drug resistant variants. Drugs are chosen on the basis of susceptibility of the test organism, mutual compatibility [some in vivo drug antagonisms are known (Tanaka, 1975)], stability at 37°C, minimal toxicity to cell cultures, and mycoplasmacidal activity, if possible. For other antibiotic susceptibility testing methods, see Section C in this volume.
Treatment Protocol 1. Cells to be cured are harvested and resuspended in the treatment cocktail to a density usually four times as high as that of the original culture. The treatment cocktail consists of the recommended growth medium for the cell line, the recommended serum, the two antibiotics of choice at a level below cellular toxicity, and additional growth factors, if necessary. Immune serum against the mycoplasma contaminant may be added to a final concentration of 5%. 2. Twofold dilutions of the cell suspension are performed in treatment cocktail; cell suspensions are diluted from 1:2 to 1:32 or 1:128, depending on the plating efficiency of the cell line. These dilutions are planted in 25-cm2 flasks. After 24 hours the cells are fed the treatment cocktail. Exposure continues for an additional 24-72 hours, depending on the microscopic assessment of cytotoxicity. 3. Following treatment, cells are either refed with or subcultured into standard growth medium with no antiserum and drugs. Generally, the lower initial planting dilutions are subcultured at this time. 4. Routine procedures for subculturing and feeding of the cells are resumed. 5. Cells are monitored for mycoplasmas at each subsequent passage and are judged free of mycoplasmas after a minimum of three consecutive negative tests.
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Discussion Two elements of this cure protocol should be emphasized. We urge pretesting of antibiotics with pure cultures of individual mycoplasma strains to ensure efficacy. We recommend dilution of the cell population in the presence of the antibiotics rather than simple addition of the drugs to a suspension or a monolayer. Once cultures are free of contamination, seed stocks should be prepared and frozen in liquid nitrogen. The frozen stocks can be used if working cultures become contaminated again. The cured cell cultures, for that matter all cell cultures, should be monitored weekly to ensure that cultures are not reinfected.
References Barile, M. F. (1979). Mycoplasma—tissue cell interactions. "/« "The Mycoplasmas" (J. G. TuUy and R. F. Whitcomb, eds.), Vol. 2, pp. 457-459. Academic Press, New York. Del Giudice, R. A., and Gardella, R. S. (1984). Mycoplasma infection of cell culture: Effects, incidence, and detection. In "Use and Standardization of Vertebrate Cell Cultures" (M. K. Patterson, ed.) In Vitro Monogr. No. 5, pp. 104-115. Tissue Culture Assoc, Gaithersburg, MD. Del Giudice, R. A., Robillard, N. F., and Carski, T. R. (1976). Immunofluorescence identification of mycoplasma on agar by use of incident illumination. J. Bacteriol. 93, 1205-1209. Drexler, H. G., Gignac, S. M., Hu, Z., Hopert, A., Fleckenstein, E., Voges, M., and Uphoff, C. C. (1994). Treatment of mycoplasma contamination in a large panel of cell cultures. In Vitro Cell Dev. Biol. 30A, 344-347. Fleckenstein, E., Uphoff, C. C , and Drexler, H. G. (1994). Effective treatment of mycoplasma contamination in cell lines with Enrofloxacin (Baytril). Leukemia 8, 1424-1434. Gardella, R. S., and Del Giudice, R. A. (1984). Antibiotic sensitivities and elimination of mycoplasmas from infected cell cultures. Isr. J. Med. Sci. 20, 931-934. Gardella, R. S., and Del Giudice, R. A. (1995). Growth of Mycoplasma hyorhinis cultivar a on semisynthetic medium. Appl. Environ. Microbiol. 61, 1976-1979. Marcus, M., Lavi, U., Nattenberg, A., Rottem, S., and Markowitz, O. (1980). Selective killing of mycoplasmas from contaminated mammalian cells in cell cultures. Nature (London) 285, 659660. Mowles, J. M. (1988). The use of ciprofloxacin for the elimination of mycoplasma from naturally infected cell lines. Cytotechnology 1, 355-358. Tanaka, N. (1975). Aminoglycoside antibiotics. In "Antibiotics" (J. W. Corcoran and F. E. Hahn, eds.). Vol. 3, pp. 345-353. Springer-Verlag, New York. Van Diggelen, O. P., Shin, S., and Phillips, D. M. (1977). Reduction in cellular tumorigenicity after mycoplasma infection and elimination of mycoplasmas from infected cultures by passage in nude mice. Cancer Res. 37, 2680-2687.
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F6 PREVENTION AND CONTROL OF MYCOPLASMA INFECTION OF CELL CULTURES Ann Smith and Jon Mowles
Introduction Mycoplasmas have a wide range of effects on their vertebrate or invertebrate cell hosts; these include effects on growth and morphology of cells, cell metabolism, and on viruses or other microbial agents cultivated in cell cultures (McGarrity and Kotani, 1985). It is therefore essential that cell lines used for research or the production of biologies be free of these contaminants. Also, a strong case can be made that some minimum standards should be imposed on the authors of scientific papers to define the mycoplasma status of any cell lines employed (Mowles and Doyle, 1990). This chapter covers laboratory procedures used to prevent and control mycoplasma contamination of cell cultures.
Source of Cells Avoid introducing mycoplasma-infected cell lines into the laboratory. Whenever possible, obtain cells from reputable culture collections, such as the American Type Culture Collection (ATCC, Rockville, MD) or the European Collection of Animal Cell Cultures (ECACC, CAMR, Salisbury, Wiltshire SP4 OJG, UK). These culture collections came about in response to the need for well445 Molecular and Diagnostic Procedures in Mycoplasmology, Vol. II
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characterized, contaminant-free cell culture seed stocks. While the rationale for development and use of established collections is well recognized by many investigators, poorly characterized cell stocks are still exchanged all too frequently.
Quarantine It is essential that once a new cell culture arrives in a laboratory there are defined procedures for handling it. It is particularly important that cultures should be handled in a Class 2 safety cabinet which offers operator protection since the source of the cell lines may not be known. Ideally a separate laboratory should be used for new cell lines (Scheirer, 1987). If this is impossible, new lines should be handled at the end of the working day, thus ensuring that clean lines remain free of mycoplasma infection. All new cell lines should be tested for mycoplasma irrespective of their source. As soon as possible, a sample of the cell line should be frozen to provide a resource in case the cell line is found to be clean and later becomes infected with mycoplasmas. Once the status of the new line has been established, the clean culture may be transferred to the main tissue culture laboratory and further repository and working cell lines prepared.
Cell Banking Once the line has been shown to be free of mycoplasma, a master cell bank should be produced. It is on this master cell bank that the major quality control tests (bacteria, fungi, mycoplasmas, and viruses) and authentication tests (karyology, isoenzyme analysis, and DNA fingerprinting) are performed. Ampoules from this bank are used to provide the working bank. When the working bank is depleted of a certain cell line, a fresh one is made from an ampoule in the master bank. The level of quality control and authentication performed on each bank depends on the end use of the line. It is, however, recommended that as a minimum both banks should be tested for bacteria, fungi, and mycoplasmas and the cell line species be verified (for more information on banking and associated testing see Bolton et al, 1993).
Sources of Mycoplasma Infection Without doubt the major source of mycoplasma contamination of cell cultures today is from other infected cultures. It has been our experience that in some
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laboratories every cell line in that laboratory, no matter what its original source, will become infected with the same mycoplasma strain due to poor laboratory management and/or poor staff training. This, of course, does not explain the origin of infection. The first large-scale systematic studies of mycoplasma contamination of cell cultures (Del Giudice and Hopps, 1978; Del Giudice and Gardella, 1984) suggested that mycoplasmas isolated from cell cultures can be divided into three main groups. Mycoplasmas from human sources were the most prevalent group and accounted for approximately one-third of all strains. Mycoplasma orale, an organism present in the human oral cavity of many healthy adults, was the single most common isolate, and strains of M. salivarium, another commensal from the human throat, have also been identified in cells. M. fermentans, an earlier infrequent contaminant, is now being reported much more frequently in cell infections. This organism is a known inhabitant of the human urogenital tract, as well as in other human organs and tissues (see Chapter D5, this volume). A possible explanation for the more numerous occurrences of this organism in cell infections comes from the increased use of primary human tissue or peripheral blood cells (lymphocytes, macrophages, etc.) as cell culture systems. The same situation may apply to M. pirum. This organism is a relatively rare cell culture contaminant, but until recently, the host origin of the organism has been unknown. Several isolates of M. pirum have been recovered from human lymphocytic cell lines (see also Chapter D5, this volume), again suggesting a human origin. In general, however, original tissue samples that are used to produce continuous cell lines are rarely contaminated with mycoplasmas. Mycoplasmas of bovine origin account for another third of all strains isolated. These include M. arginini, M. bovis, and Acholeplasma laidlawii. Commercial bovine serum is the source of these species, and improvements in filter technology and other commercial efforts to collect serum in a more clean environment and within a closed collection system should help reduce the chances of mycoplasma contamination of commercial bovine serum. All other mycoplasmas make up the final third of the strains isolated. By far the most common of this group is M. hyorhinis, a species of swine origin. Trypsin has been suggested as the source of M. hyorhinis because trypsin is obtained from the hog pancreas. However, the shortcoming of this argument is the lack of supporting data; despite many attempts in many laboratories, mycoplasmas have never been isolated from trypsin preparations. Further, even if mycoplasmas were originally present in the pancreas, they are unlikely to survive the purification scheme used in commercial trypsin preparation. Thus, there is no demonstrable explanation for the widespread occurrence of M. hyorhinis in cell cultures, and there are no confirmed reports of finding M. hyorhinis in bovine serum. Undoubtedly, cell to cell transfer accounts for the high proportion of the strains of M. hyorhinis isolated through these years. Over an 8-year period at the European Collection of Animal Cell Cultures the most common cell culture contaminants were M. arginini (42%), M. orale
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(29%), and M. fermentans (22%). During the same period the percentage of positive lines of those cell lines tested ranged from 2 to 22% per year. However, the reported incidence of infected cell lines in various geographic locations has varied considerably and in some cases may exceed 50% (McGarrity and Kotani, 1985).
Control of Infection When testing cell lines for mycoplasmas it is important that more than one detection method be used since all methods have their limitations. Cultivation on agar is the most sensitive and specific method to detect mycoplasmas but it is necessary to understand that cultivar a strains of M. hyorhinis will not grow on any of the conventional mycoplasma media currently described. For methods to detect M. hyorhinis, which does not rely on broth or agar cultivation, see Chapters F3 and F4 in this volume. It should also be noted that the level of sensitivity and specificity of the available commercial detection kits varies considerably and this must be kept in mind when selecting and using such kits. For example, a kit that detects only high levels of infection or only certain Mycoplasma or Acholeplasma species is of little value. The use of an inadequate test may be worse than not testing at all because it can lead to a false sense of security. Prior to testing, cells should undergo at least two passages in antibiotic-free media and also two passages from frozen resuscitation since antibiotics and cryoprotectants can have an adverse effect on the growth of mycoplasmas (possibly lowering their levels to below the limit of detection). The frequency with which cells should be tested is dependent on the type of laboratory and the volume of cell cultures employed. Laboratories receiving many cell lines should test more frequently than those that culture only a few lines. Lines should always be tested on initial receipt, on resuscitation, and before freezing to ensure a mycoplasma-free stock of cells. As noted earlier, testing should preferably be carried out in a location quarantined from other cell production facilities. If specific antimycoplasma antibiotics have been used in an attempt to cure a culture of infection, testing over a period of five consecutive passages in antibiotic-free media should be made before the culture can confidently be called mycoplasma negative. Once a culture is found to be mycoplasma positive, immediate action is necessary to avoid subsequent infection of other lines within the same laboratory. All media and other reagents associated with the infected line should either be discarded or checked for the presence of mycoplasmas. To have confidence in the removal of the source of the contaminant it is better, under most circumstances, to discard the cell line.
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Ideally, a thorough disinfection of all work surfaces and safety cabinets should follow. Other cell lines growing within the same laboratory should be immediately checked for contamination and regarded as potentially contaminated until proven otherwise. By using the just-mentioned procedure the chances of secondary infection of other lines are reduced to a minimum. Suppliers of bovine serum do mycoplasma testing on their product, although procedures and quality control of medium components are not well standardized. However, mycoplasmas present in low titers and limited lot sampling can contribute to problems in detection. The larger the volume tested, the greater the confidence level in negative results. If additional tests are to be performed in a working cell culture laboratory, the large-volume serum test method is recommended (see Chapter F2, this volume) and should incorporate as many samples as is economically feasible. The best method of treatment of serum is heat inactivation at 56°C for 1 hour, in volumes no larger than 100 ml. A preliminary test should be done to determine whether the heated serum will support the growth of the particular cell in use, using the same unheated serum as a control. However, most cells will grow well with heat-inactivated serum. Antibiotics should never be relied on to prevent mycoplasma infection. The unfortunate practice of maintaining cell lines on antibiotic-containing media is probably the single most important factor responsible for the lateral spread of mycoplasma contamination of cell cultures. There are some specific applications where antibiotics may be useful, such as in attempts to isolate viruses from frankly contaminated clinical material or in procedures to establish a new cell line from contaminated tissue. However, there is absolutely no justification for the use of antibiotics in the routine maintenance of cell cultures. When a break in aseptic technique occurs in the absence of antibiotics, it is usually macroscopically apparent by microbial overgrowth in the nutrient-rich cell culture system. When this happens, the contaminated cultures are discarded and there is no further amplification of the problem. Antibiotics that are commonly used in cell culture media such as penicillin and streptomycin can mask a break in aseptic technique by preventing microbial outgrowth. However, these antibiotics are ineffective against mycoplasmas so any mycoplasma contamination introduced through faulty asepsis is propagated with each cell culture passage. This amplifies the contamination problem and places other cultures at risk. It should be obvious that reliance on antibiotics belies inadequate aseptic technique.
Discussion An awareness of sources of infection and a program of cell maintenance designed to reduce risk of exposure of cells to these sources are of principal
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importance in preventing mycoplasma infection of cell cultures. As indicated in previous sections, human sources and bovine serum make up two of the main reservoirs. The lateral spread of mycoplasmas from one cell line to another is a more important factor today than the original source because the infected cells become a reservoir from which infection is spread to other cells. Mycoplasmas of human origin continue to be the predominant contaminants. Strict adherence to aseptic technique is essential to reduce exposure to these strains. With the exchange of cultures between laboratories and as the number of cell lines within each laboratory increases, the opportunity for mycoplasma transfer between cell lines increases. Cell lines untested or positive for mycoplasmas should be quarantined. Mycoplasma-free cultures should be segregated from infected cultures by time or place of handling. Reagents for the two sets of cells should be separate. Occasionally, mycoplasma spread between cell lines has been attributed to the use of a contaminated virus pool. These should be tested for mycoplasmas routinely. Some generalizations are appropriate for successful cell handling. A program of routine monitoring for mycoplasmas should be adopted, and techniques to identify the Mycoplasma or Acholeplasma species involved may help focus on the source of the contamination. The use of antibiotics is not recommended, except in situations where primary tissues or large-volume culturing is undertaken. Use of antibiotics may lead to lapses in aseptic technique, to selection of drug-resistant organisms, and to delayed detection of low-level infection by either mycoplasmas or other bacteria. Stocks of mycoplasma-free cell lines should be frozen and stored to provide a continuous supply of cells should working stocks become contaminated. Perhaps the most important thing to remember is that one need not go to extraordinary lengths to prevent mycoplasma contamination. The vast majority of cell cultures are free of mycoplasmas and they remain free even after continuous passage under commonplace laboratory conditions. Common sense and sound aseptic techniques are all that is required.
References Bolton, B. I., Morris, C. B., and Mowles, J. M. (1993). Long term cultures and cell lines. In "Methods of Immunological Analysis" (R. F. Masseyeff, W. H. Albert, and N. A. Staines, eds.). Vol. 3, pp. 103-120. VCH, Weinheim. Del Giudice, R. A., and Gardella, R. S. (1984). Mycoplasma infection of cell culture: Effects, incidence, and detection. In "Use and Standardization of Vertebrate Cell Cultures" (M. K. Patterson, ed.) In Vitro Monogr. No. 5, pp. 104-115. Tissue Culture Assoc, Gaithersburg, MD.
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Del Giudice, R. A., and Hopps, H. E. (1978). Microbiological methods and fluorescent microscopy for the direct demonstration of mycoplasma infection of cell cultures. In "Mycoplasma Infection of Cell Cultures" (G. J. McGarrity, D. G. Murphy, and W. W. Nichols, eds.), pp. 57-69. Plenum, New York. McGarrity, G. J., and Kotani, H. (1985). Cell culture mycoplasmas. In "The Mycoplasmas" (S. Razin and M. F. Barile, eds.). Vol. 4, pp. 353-390. Academic Press, Orlando, FL. Mowles, I. M., and Doyle, A. (1990). Cell culture standards—time for a rethink? Cytotechnology 3, 107-108. Scheirer, W. (1987). Laboratory management of animal cell culture processes. Trends Biotechnol. 5, 261-265.
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Appendix
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APPENDIX
TABLE I GENUS Mycoplasma AND MAJOR CHARACTERISTICS
Species and Type Strain^' Mycoplasma adleri G-\45'^ Mycoplasma agalactiae PG2T Mycoplasma alkalescens T>\2^ Mycoplasma alvi Ilsley'^ Mycoplasma anatis 1340^ Mycoplasma anseris 1219^^ Mycoplasma arginini G230'r Mycoplasma arthritidis PG6T Mycoplasma auris UlA'^ Mycoplasma bovigenitalium PGW^ Mycoplasma bovirhinis PG43'^ Mycoplasma bovis Donetta"^ Mycoplasma bovoculi M165/69T Mycoplasma buccale CH20247T Mycoplasma buteonis BbT2g'r Mycoplasma californicum ST-6^ Mycoplasma canadense 275C^ Mycoplasma canis ?GW Mycoplasma capricolum subsp. capricolum Calif. Kid^ subsp. capripneumoniae F38'^
DNA base composition (G + C mol%)
Biochemical properties glucose/arginine^
29.6 30.5-34.2 25.9 26.4 26.6 27.4-26.0 27.6-28.6 30.0-32.6 26.9 28.1-30.4 24.5-27.3 27.8-32.9 29.0 25.0-26.4 27.0 31.9 29.0 28.4-29.1
-/ + -/-/ + +/+ +/— —/+ —/+ -/+ -/ + —/+/— —/— +/-/+ +/-/-/+ +/-
24.1-25.5 24.4
+/+ +/-
Principal host(s) Caprine Caprine/ovine Bovine Bovine Avian Avian Caprine/ovine Murine Caprine Bovine Bovine Bovine Bovine Human/primate Avian Bovine Bovine Canine/bovine Caprine Caprine {continues) 455
456
Appendix
TABLE I
Species and Type Strain" Mycoplasma caviae 0122"^ Mycoplasma cavipharyngis WIC^ Mycoplasma cite Hi RG-2CT Mycoplasma cloacale 383"^ Mycoplasma collis 586'^ Mycoplasma columbinasale 694"^ Mycoplasma columbinum MMP-l^ Mycoplasma columborale MMP-4'r Mycoplasma conjunctivae HRC58F Mycoplasma corogypis B V F Mycoplasma cottewii VIS'^ Mycoplasma cricetuli CW^ Mycoplasma cynos H831^ Mycoplasma dispar 462/2^ Mycoplasma edwardii PG24'r Mycoplasma equigenitalium T31^ Mycoplasma equirhinis M432/72'^ Mycoplasma falconis H/Tl"^ Mycoplasma fastidiosum 4822'^ Mycoplasma faucium DC333'r Mycoplasma felifaucium PU'^ Mycoplasma feliminutum BQU^ Mycoplasma felis CO'^ Mycoplasma fermentans PGIK^ Mycoplasma flocculare MS42T Mycoplasma gallinaceum DD'^ Mycoplasma gallinarum PG16T Mycoplasma gallisepticum VG?>Y^ Mycoplasma gallopavonis WRl'^ Mycoplasma gateae CS"^ Mycoplasma genitalium G37'r Mycoplasma glycophilum 486"^ Mycoplasma gypisBXITY^ Mycoplasma hominis PG2F Mycoplasma hyopharyngis H3-6BFT Mycoplasma hyopneumoniae F Mycoplasma hyorhinis BTS7'r Mycoplasma hyosynoviae S\6^ Mycoplasma imitans 4229'^ Mycoplasma indiense 3T^ Mycoplasma iners PGSa^ Mycoplasma iowae 695"^
(continued)
DNA base composition (G + C mol%)
Biochemical properties glucose/arginine*
Principal host(s)
nd^ 30.0 27.4 26.0 28.0 32.0 27.3 29.2 nd 28.0 27.0 nd 25.8 28.5-29.3 29.2 31.5 nd 27.5 32.3 nd 31.0 29.1 25.2 27.5-28.7 33.0 28.0 26.5-28.0 31.8-35.7 27.0 28.5 32.4 27.5 21A 27.3-33.7 24.0 27.5-33.0 27.3-27.8 28.0 31.9 32.0 29.1-29.6 25.0
+/+/+/—/+ +/-/+ -/+ +/— +/+/+/— +/— +/— +/+/+/—/+ —/+ +/— -/+ —/+ +/— +/— +/+ —/— +/— -/+ +/— +/— -/+ +/+/— -/+ -/+ -/+ -/+1-/+ +/— —/+ -/+ +/+
Rodent/guinea pig Rodent/guinea pig Rodent/squirrel Avian Canine Avian Avian Avian Ovine/bovine Avian Caprine Rodent/hamster Canine Bovine Canine Equine Equine Avian Equine Human/primate Feline Feline Feline/equine Human/primate Porcine Avian Avian Avian Avian Feline Human Avian Avian Human Porcine Porcine Porcine Porcine Avian Primate Avian Avian (continues)
457
Appendix
TABLE I
Species and Type Strain^ Mycoplasma leocaptivus 3L2'r Mycoplasma leopharyngis LL2^ Mycoplasma lipofaciens R 1 7 F Mycoplasma lipophilum MaBy'^ Mycoplasma maculosum PG15'^ Mycoplasma meleagridis 17529"^ Mycoplasma moatsii MK405'r Mycoplasma mobile 163K'^ Mycoplasma molare H542'^ Mycoplasma muris RIIM"^ Mycoplasma mustelae MX9'^ Mycoplasma mycoides subsp. capri PG3'r subsp. mycoides P G F Mycoplasma neurolyticum Type A'^ Mycoplasma opalescens MH5408'^ Mycoplasma ovale CH19299T Mycoplasma ovipneumoniae ¥98'^ Mycoplasma oxoniensis 128"^ Mycoplasma penetrans GTU54'r Mycoplasma phocacerebrale 1049'^ Mycoplasma phocarhinis 852"^ Mycoplasma phocidae 105'^ Mycoplasma pirum 70-159"^ Mycoplasma pneumoniae FH^ Mycoplasma primatum HRC292T Mycoplasma pullorum CKK'^ Mycoplasma pulmonis PG34'^ Mycoplasma putrefaciens KS-1''' Mycoplasma salivarium PG20'r Mycoplasma simbae LX"^ Mycoplasma spermatophilum AH 159'^ Mycoplasma spumans PG13'r Mycoplasma sualvi Mayfield B'^ Mycoplasma subdolum TB'^ Mycoplasma synoviae WVU 1853"^ Mycoplasma testudinis 01008"^ Mycoplasma yeatsii GUU Mycoplasma verecundum 107^
(continued)
DNA base composition (G + C mol%)
Biochemical properties glucose/arginine^
27.0 28.0 24.5 29.7 26.7-29.6 27.0-28.6 25.7 23.5 26.0 24.9 28.2
+/-/+/+ -/+ -/+ -/+ +/+ +/+/-/+ +/-
Feline Feline Avian Human/primate Canine Avian Primate Piscine Canine Rodent/mice Rodent/mink
24.0-26.0 26.1-27.1 22.8-26.2 29.2 24.0-28.2 25.7 29.0 30.5 25.9 26.5 27.8 25.5 38.6-40.8 28.6 29.0 27.5-29.2 28.9 27.3-31.4 37.0 32.0 28.4-29.1 23.7 28.8 34.2 35.0 26.6 27.0-29.2
+/+/+/-/+ -/+ +/+/+/-/+ -/+ -/+ +/+ +/-/+ +/+/+/-/+ -/+ -/+ -/+ +/+ -/+ +/+/+/+ -/-
Caprine Bovine/caprine Murine Canine Human/primate Ovine Rodent/hamster Human Aquatic/seal Aquatic/seal Aquatic/seal Human Human Primates/human Avian Murine Caprine Human Primate Human Canine Porcine Equine Avian Reptile/turtle Caprine Bovine
Principal host(s)
«Superscript T denotes type strain. ^Positive or negative responses in glucose fermentation/arginine hydrolysis tests. ^Not done.
458
Appendix
TABLE II GENERA UREAPLASMA AND ACHOLEPLASMA
Species and Type Strain^ Ureaplasma canigenitalium D6P-C^ Ureaplasma cati ¥2^ Ureaplasma diversum A417'r Ureaplasma felinum FT2-B'^ Ureaplasma gallorale D6-l^ Ureaplasma urealyticum T960T^ Acholeplasma axanthum S-743'^ Acholeplasma brassicae 0502"^ Acholeplasma cavigenitalium G?V^ Acholeplasma equifetale 0112"^ Acholeplasma granularum BTS39'r Acholeplasma hippikon C P Acholeplasma laidlawii PG8^ Acholeplasma modicum PG49'^ Acholeplasma morum 72-043'^ Acholeplasma multilocale PN525''' Acholeplasma oculi IQL'^ Acholeplasma palmae 1-233'^ Acholeplasma parvum H23M^
DNA base composition (G + C mol%)
Principal host(s)
28.7 28.1 26.9-27.8 27.1 27.6 26.9-27.8 31.0 35.5 36.0 30.5 30.0-32.0 33.1 31.0-36.0 29.0 34.0 31.0 26.0-27.0 30.0 29.1
Canine Feline Bovine Feline Avian Human Bovine/porcine/plants Plants Rodent/guinea pig Equine Porcine/caprine/ovine Equine Many animals/plants Bovine/equine Bovine Equine Capine/equine Plants Equine
'^All Ureaplasma species hydrolyze urea and do not ferment glucose or hydrolyze arginine. All Acholeplasma species, except A. parvum, ferment glucose and do not hydrolyze arginine or urea.
TABLE III GENERA ANAEROPLASMA AND ASTEROLEPLASMA
Species and Type Strain^
Base composition (G + C mol%)
Anaeroplasma ahactoclasticum 6-n Anaeroplasma bactoclasticum }K^ Anaeroplasma intermedium IhAJ Anaeroplasma varium k-2^ Asteroleplasma anaerobium 161^
29.3-30.1 32.8-33.7 32.5 33.4 39.2-40.5
'^AU species are strictly anaerobic and oxygen-sensitive, and occur so far only in the bovine or ovine rumen.
459
Appendix
TABLE IV GENERA ENTOMOPLASMA AND MESOPLASMA
Species and Type Strain"
DNA base composition (G + C mol%)
Entomoplasma ellychniae ELCN-l"^ Entomoplasma lucivorax PIPN-2'^ Entomoplasma luminosum PIMN-F Entomoplasma melaleucae Ml'^ Entomoplasma somnilux PYAN-l^ Mesoplasma chauliocola CHPA-2'^ Mesoplasma coleopterae BARC-779'^ Mesoplasma corruscae EIXA-2^ Mesoplasma entomophilum TAC^
27.5 27.4 28.8 27.0 27.4 28.3 27.8 26.5 30.0
Mesoplasma florum Ll^ Mesoplasma grammopteriae GRUA-F Mesoplasma lactucae 831-C4'^ Mesoplasma photuris PUPA-2'^ Mesoplasma pleciae PS-1^ Mesoplasma seiffertii FT'^ Mesoplasma syrphidae YIS"^ Mesoplasma tabanidae BARC-SS?"^
27.3 29.1 30.0 29.8 31.6 30.0 27.6 28.1
Principal host(s) Firefly Firefly Firefly Plant/bee Firefly larva Beetles Beetles/plants Firefly Beetles, horseflies, bees, moths, butterflies, plants Plants/beetles Beetles/bee Plant Fireflies Fly maggot Plants/deerfly/mosquitoes Flies/bee/butterfly Horsefly
"All species ferment glucose and, with exception oi Me. photuris, do not hydrolyze arginine.
TABLE V GROUPCLASSIFICATION OF GENUSSpiroplasma
Binomial andlor common name Spiroplasma citri
S. kunkelii
277F spiroplasma Green leaf bug S. insolitum Cocos spiroplasma S. phoeniceum
Sex ratio spiroplasmas S. floricola
Groupa
Strainsb Maroc-R8A2T(27556) C 189(27665) Israel BC-3T(33219) AS 576(29416) E275T(29320) I-747(2905 1) B655(33289) 277F(29761) LB- 12(33649) MST(33502) N525(33287) N628 P40T(43115)
23-6T(29989) BNR l(33220) OBMG(33221)
G+C content (mol%)
Glucose/ arginine
Principal host(s)
Disease incited
Dicots, leafhoppers
Citrus stubborn
Bees Maize, leafhoppers
Honeybee spiroplasmosis Corn stunt
Rabbit tick Green leaf bug Flowers, Ersitalis fly Coconut palm
None None None None
Catharanthus roseus Drosophila
Periwinkle disease Sex ratio trait
Insects. flowers
None known
known known known known
S . apis
IV
S . mirum
V
S. ixodetis S. monobiae Syrphid spiroplasma
VI VII
EA- l(33826) DF- l(43209) TAAS-l(5 1123) CN-ST(33827) AES- 1T(35112) MQ-4T(35262) DU-l(43210)
S . clarkii S . culicicola S. velocicresens Cucumber beetle S. sabaudiense
Ellychnia Leafhopper S. cantharicola
Vacant.Tabanid spiroplasma
B31T(33834) SR 3(33095) PPS l(33450) SMCAT(29335) GT-48(29334) TP-2(33503) Y32T(33835) MQ- lT(33825)
XI11 XIV
xv XVI-I XVI-2 XVI-3 XVII XVIII
Ar- 1343T(43303) EC- l(432 12) I-25(43262) CC- lT(43207) CB-l(43208) AR- 1357(5 1126)
30
+I+
Bees, flowers
"May disease"
30
+/+
Rabbit ticks
25
+/-
Ixodes pacificus ticks Monobia wasp
Suckling mouse cataract disease None known None known
Eristalis arbustorum fly Chrysops deerfly Tabanus atratus horsefly Cotinus beetle Aedes mosquito Monobia wasp Diabrotica undecimpunctata beetle Aedes mosquito Ellychnia corrusca beetle Cicadulina leafhopper Cantharis beetle beetle mosquito
None None None None None None None
known known known known known known known
None None None None
known known known known
Tabanus nigrovittatus horsefly
None known
(continues)
TABLE V
Binomial and/or common name
Groupa
Firefly spiroplasma
XIX
PUP- l(43206)
Colorado potato beetle spiroplasma Flower spiroplasma S. taiwanense
XX
LD- l(432 13)
XXI XXII
W 115(43260) CT- lT(433O2)
Tabanid spiroplasma
XXIII
S. chinense
XXIV
Strainsb
S. diminutum
XXV Tentative group designationsd (XXVI) (XXVII) (XXVIII) (XXIX) (XXX) (XXXI) (XXXII) (XXXIII) -
-
groups
PLHS-l(5 1752) TALS-2(5 1749) PALS- l(5 1748) TIUS-l(51751) BIUS-l(51750) HYOS- 1(5 1745) TABS-2(5 1746) TAUS- 1(5 1747)
(continued)
G+C content (mol%)
Glucose/ arginine
Principal host(s) Photuris pennsylvanicus beetle Leptinotarsa decemlineata beetle Prunus sp. flower Culex tritaeniorhynchus mosquito Tabanus gladiator horsefly Calystegia hederaceae flowers Culex annulus mosquito
Disease incited None known None known
None known None known None known None known None known
Scorpionfly Horsefly Dragonfly Tiphiid wasp Flower Horsefly Horsefly Horsefly
-
assigned on the basis of failure to cross-react in growth inhibition, metabolism inhibition, and deformation serological
tests. bAccession numbers (in parentheses) from American Type Culture Collection. Strain DF- I previously assigned this group designation has been reclassified to Group VIII-2. "Proposed group designations in manuscript in preparation.
Index
Acholeplasma species, and host associations, 458 Acquisition, mollicutes, 2-3 AIDS ELISA for mollicute detection, 115- 121 laboratory diagnostic tests for mollicutes, 247-254 Anaeroplasma species, and host associations, 458 Animals, mollicute species in, 9-12, 455-458 Antibiotic sensitivity tests, 181-204 Antibiotics cidal activity testing, 199-204 determination of minimum inhibitory concentration, 189-1 97 use in treatment of mollicute-infected cell cultures, 439-443 Aquatic hosts, mollicutes in, 11-12 Arthritis, experimental models, 349-359 Arthropod mollicutes, 12- 14, 459-462 Asteroleplasma species, and host associations, 458 Avian mollicutes, 11, 283-292, 361-370 experimental infections with, 361-370 Axenic cultivation, cell culture mollicutes, 411-418
Bovine mollicutes, 9-10, 255-264 Bovines diagnostic tests for mollicutes, 255-264 experimental mollicute infections, 377-384
Canine mollicutes, 10- 11 Caprine mollicutes, 10, 265-273 Cell cultures mollicute detection by axenic cultivation, 41 1-418 by PCR,431-438 by staining techniques, 419-429
mollicute infections, 407-451 prevention and control, 443-451 treated with antibiotics, 439-443 Cultivation, mollicutes, from cell culture infections, 411-418
Diagnosis cell culture mollicute infections, 407-451 mollicute infections, use of monoclonal antibodies in, 137- 146 Diagnostic tests avian mollicutes, 283-292 bovine mollicutes, 255-264 caprine mollicutes, 265-273 mollicutes, 207-322 mollicutes in AIDS patients, 247-254 mollicutes in insects, 313-322 Mycoplasma pneumoniae, 2 11-223 neonatal infections, 237-245 ovine mollicutes, 265-273 porcine mollicutes, 275-28 1 sexually transmitted diseases, 225-236 spiroplasma infections in plantslinsects, 293-297 DNA amplification and product identification, 65-73 DNA fragments, in mollicute detection and identification, 47-5 1 DNA preparation, for PCR analysis, 61-64 DNA staining, for mollicute detection in cell cultures, 419-429 rDNA probes, for mollicute detection and phylogeny, 29-46
E Ecology, mollicutes, 7- 15 ELISA in avian hosts, 93- 104 in bovine hosts, 105-1 13, 137-146 in caprine hosts, 105-1 13 in human respiratory infections, 123-136
464
Index
in human urogenital infections and AIDS, 115-121 in laboratory animal rodents, 93-104 in ovine hosts, 105-113 in phytoplasma (MLO) infections, 141-146 in porcine hosts, 105-113 in small animal hosts, 93-104 Entomoplasma species, and host associations, 459 Equine mollicutes, 10 Experimental infections cattle, 377-384 cell cultures, 399-403 plants/insects, 385-389, 391-398 poultry, 361-370 swine, 371-376 Experimental moUicute infections, 325-403 Experimental respiratory infections, in rodents, 327-336 Experimental urogenital infections, in rodents, 337-347
Feline mollicutes, 10-11 Fluorescence, for detection of mollicutes in cell cultures, 419-429
H Habitat, mollicutes, 7-15 role in laboratory diagnostic techniques, 1517 HIV/mycoplasma infections, in cell cultures, 399-403 Hosts colonization with mollicutes, 3-4 infection with mollicutes, 3-4 specificity of mollicutes for, 5-7 Humans, mollicute species in, 7-9, 455-458
I Immunoblotting/immunobinding, in mollicute antigen/antibody analysis, 151-167 Immunofluorescence, in diagnosis of mollicute infections, 147-150, 169-177, 419-429 Insect (arthropod) mollicutes, 12-14, 459-462
Insects identification of mollicutes in, 313-322 laboratory techniques for spiroplasmas in, 293-297
Laboratory animal mollicutes, 11, 93-104, 327-347 Laboratory diagnosis, mollicutes, habitat considerations, 15-17 M Mesoplasma species, and host associations, 459 Microimmunofluorescence, in mollicute antibody detection, 147-150 Mollicute detection 16S rRNA oligonucleotide probes in, 29-46 by nested PCR, 75-79 cloned genomic DNA fragments in, 47-51 in cell culture infections by PCR, 431-438 in cell cultures with staining techniques, 419-429 Mollicute infections, diagnosis by serologic methods, 89-177 Mollicutes acquisition and transmission of, 2-3 antibiotic sensitivity, 181-204 antibody detection by immunoblotting/immunobinding, 151167 by microimmunofluorescence, 147-150 cultivation from cell culture infections, 4 1 1 418 current classification of genus and species, 455-462 diagnostic tests for, 207-322 experimental arthritis models with, 349-359 experimental infections, 325-403 experimental poultry infections, 361-370 experimental rodent respiratory infections, 327-336 experimental rodent urogenital infections, 337-347 habitat and role in laboratory diagnoses, 1517 host colonization versus infection, 3-4
Index host interrelationships of, 1-21, 455-462 in animals, 9-12, 255-281 in aquatic hosts, 11-12 in arthropods (including insects), 12-14 in avian hosts, 11, 283-292 in bovines, 9-10, 255-264 in canines, 10-11 in caprines, 10, 265-273 in cell culture infections, 407-451 in equines, 10 in felines, 10-11 in humans and nonhuman primates, 7-9, 211-254 in insects, 12-14 in laboratory rodents, 11 in large domestic animals, 9-12 in ovines, 10, 265-273 in plants, 14-15 in porcines, 10, 275-281 in wild animals, 11-12 prevention and control of cell culture infections, 443-451 serological methods in detection and diagnoses, 89-177 strain comparison by arbitrary primer PCR, 81-85 treatment of cell culture infections with antibiotics, 439-443 Monoclonal antibodies, as diagnostic tools, 137-146 Mycoplasma genitalium, detection by immunofluorescence, 169-177 Mycoplasma pneumoniae detection by immunofluorescence, 169-177 laboratory diagnosis, 211-223 Mycoplasma species, and host associations, 455-457 Mycoplasmacidal activity, antibiotics, tests for, 199-204 Mycoplasmas/HIV cell culture infections, 399-403 N Neonatal infections, diagnosis, 237-245 O Ovine mollicutes, 10, 265-273
465
PCR amplification and identification of products, 65-73 moUicute detection in cell culture infections, 431-438 nested PCR for mollicute detection, 75-79 preparation of DNA from clinical specimens, 61-64 random amplified polymorphic DNA fingerprinting, 81-85 selection of target sequences and primers, 53-60 Phylogeny, use of oligonucleotide probes, 2946 Phytoplasmas detection in plant tissues with ELISA, 141146 diagnostic tests in plants, 299-311 experimental plant/insect infections, 391398 Plants experimental mollicute infections, 385-389 experimental phytoplasma infections, 391398 laboratory techniques for phytoplasma detection, 299-311 laboratory techniques for spiroplasmas, 293297 mollicute infections, 14-15 phytoplasma detection by ELISA tests, 131146 Porcine mollicutes, 10, 275-281 experimental infections, 371-376 Poultry, experimental mollicute infections, 361-370 Prevention/control, mollicute infections in cell cultures, 443-451
Respiratory infections detection in man by ELISA, 123-136 experimental, in rodents, 327-336 Ribosomal DNA probes, for mollicute detection and phylogeny, 29-46 Rodents experimental respiratory infections, 327-336 experimental urogenital infections, 337-347
466
Index Transmission, mollicutes, 2-3
Serological analysis, mollicutes, 89-177 Sexually transmitted diseases, diagnostic testing for, 225-236 Spiroplasma species, and host associations, 460-462 Spiroplasmas diagnostic tests in plants/insects, 293-297 experimental plant infections, 385-389 identification techniques in insects, 313-322 Swine, experimental mollicute infections, 371376
U Ureaplasma species, and host associations, 458 Urogenital infections experimental models in rodents, 337-347 in man, detection by ELISA, 115-121 by microimmunofluorescence, 147-150
W Target sequences, selection for PCR analysis, 53-60
Wild animals, mollicutes in, 11-12
Fig. 1. Vero cells infected with M. hyorhinis, stained by the double stain method with DNAF and fluoresceinated anti-M hyorhinis. (a) and (b) show the same field, (a) Viewed with the filter set for DNAF. (b) Viewed with the filter set for immunofluorescence. The apple green color of specific fluorescein fluorescence is easily compromised in the photographic process but one should expect to see it in the microscope.
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