The Prokaryotes, Vol. 5: Proteobacteria: Alpha and Beta Subclasses

The Prokaryotes Third Edition

The Prokaryotes A Handbook on the Biology of Bacteria Third Edition

Volume 5: Proteobacteria: Alpha and Beta Subclasses

Martin Dworkin (Editor-in-Chief), Stanley Falkow, Eugene Rosenberg, Karl-Heinz Schleifer, Erko Stackebrandt (Editors)

Editor-in-Chief Professor Dr. Martin Dworkin Department of Microbiology University of Minnesota Box 196 University of Minnesota Minneapolis, MN 55455-0312 USA Editors Professor Dr. Stanley Falkow Department of Microbiology and Immunology Stanford University Medical School 299 Campus Drive, Fairchild D039 Stanford, CA 94305-5124 USA Professor Dr. Eugene Rosenberg Department of Molecular Microbiology and Biotechnology Tel Aviv University Ramat-Aviv 69978 Israel

Professor Dr. Karl-Heinz Schleifer Department of Microbiology Technical University Munich 80290 Munich Germany Professor Dr. Erko Stackebrandt DSMZ- German Collection of Microorganisms and Cell Cultures GmbH Mascheroder Weg 1b 38124 Braunschweig Germany

URLs in The Prokaryotes: Uncommon Web sites have been listed in the text. However, the following Web sites have been referred to numerous times and have been suppressed for aesthetic purposes: www.bergeys.org; www.tigr.org; dx.doi.org; www.fp.mcs.anl.gov; www.ncbi.nlm.nih.gov; www.genome.ad.jp; www.cme.msu.edu; umbbd.ahc.umn.edu; www.dmsz.de; and www.arb-home.de. The entirety of all these Web links have been maintained in the electronic version.

Library of Congress Control Number: 91017256 Volume 5 ISBN-10: 0-387-25495-1 ISBN-13: 978-0387-25495-1 e-ISBN: 0-387-30745-1 Print + e-ISBN: 0-387-33492-0 DOI: 10.1007/0-387-30745-1 Volumes 1–7 (Set) ISBN-10: 0-387-25499-4 ISBN-13: 978-0387-25499-9 e-ISBN: 0-387-30740-0 Print + e-ISBN: 0-387-33488-2 Printed on acid-free paper. © 2006 Springer Science+Business Media, LLC All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed in Singapore. 9 8 7 6 5 4 3 2 1 springer.com

(BS/KYO)

Preface

Each of the first two editions of The Prokaryotes took a bold step. The first edition, published in 1981, set out to be an encyclopedic, synoptic account of the world of the prokaryotes—a collection of monographic descriptions of the genera of bacteria.The Archaea had not yet been formalized as a group. For the second edition in 1992, the editors made the decision to organize the chapters on the basis of the molecular phylogeny championed by Carl Woese, which increasingly provided a rational, evolutionary basis for the taxonomy of the prokaryotes. In addition, the archaea had by then been recognized as a phylogenetically separate and distinguishable group of the prokaryotes. The two volumes of the first edition had by then expanded to four. The third edition was arguably the boldest step of all. We decided that the material would only be presented electronically. The advantages were obvious and persuasive. There would be essentially unlimited space. There would be no restrictions on the use of color illustrations. Film and animated descriptions could be made available. The text would be hyperlinked to external sources. Publication of chapters would be seriati—the edition would no longer have to delay publication until the last tardy author had submitted his or her chapter. Updates and modifications could be made continuously. And, most attractively, a library could place its subscribed copy on its server and make it available easily and cheaply to all in its community. One hundred and seventy chapters have thus far been presented in 16 releases over a six-year period. The virtues and advantages of the online edition have been borne out. But we failed to predict the affection that many have for holding a bound, print version of a book in their hands. Thus, this print version of the third edition shall accompany the online version. We are now four years into the 21st century. Indulge us then while we comment on the challenges, problems and opportunities for microbiology that confront us.

Moselio Schaechter has referred to the present era of microbiology as its third golden age—the era of “integrative microbiology.” Essentially all microbiologists now speak a common language. So that the boundaries that previously separated subdisciplines from each other have faded: physiology has become indistinguishable from pathogenesis; ecologists and molecular geneticists speak to each other; biochemistry is spoken by all; and—mirabile dictu!—molecular biologists are collaborating with taxonomists. But before these molecular dissections of complex processes can be effective there must be a clear view of the organism being studied. And it is our goal that these chapters in The Prokaryotes provide that opportunity. There is also yet a larger issue. Microbiology is now confronted with the need to understand increasingly complex processes. And the modus operandi that has served us so successfully for 150 years—that of the pure culture studied under standard laboratory conditions—is inadequate. We are now challenged to solve problems of multimembered populations interacting with each other and with their environment under constantly variable conditions. Carl Woese has pointed out a useful and important distinction between empirical, methodological reductionism and fundamentalist reductionism. The former has served us well; the latter stands in the way of our further understanding of complex, interacting systems. But no matter what kind of synoptic systems analysis emerges as our way of understanding host–parasite relations, ecology, or multicellular behavior, the understanding of the organism as such is sine qua non. And in that context, we are pleased to present to you the third edition of The Prokaryotes. Martin Dworkin Editor-in-Chief

Foreword

The purpose of this brief foreword is unchanged from the first edition; it is simply to make you, the reader, hungry for the scientific feast that follows. These four volumes on the prokaryotes offer an expanded scientific menu that displays the biochemical depth and remarkable physiological and morphological diversity of prokaryote life. The size of the volumes might initially discourage the unprepared mind from being attracted to the study of prokaryote life, for this landmark assemblage thoroughly documents the wealth of present knowledge. But in confronting the reader with the state of the art, the Handbook also defines where more work needs to be done on well-studied bacteria as well as on unusual or poorly studied organisms. This edition of The Prokaryotes recognizes the almost unbelievable impact that the work of Carl Woese has had in defining a phylogenetic basis for the microbial world. The concept that the ribosome is a highly conserved structure in all cells and that its nucleic acid components may serve as a convenient reference point for relating all living things is now generally accepted. At last, the phylogeny of prokaryotes has a scientific basis, and this is the first serious attempt to present a comprehensive treatise on prokaryotes along recently defined phylogenetic lines. Although evidence is incomplete for many microbial groups, these volumes make a statement that clearly illuminates the path to follow. There are basically two ways of doing research with microbes. A classical approach is first to define the phenomenon to be studied and then to select the organism accordingly. Another way is to choose a specific organism and go where it leads. The pursuit of an unusual microbe brings out the latent hunter in all of us. The intellectual challenges of the chase frequently test our ingenuity to the limit. Sometimes the quarry repeatedly escapes, but the final capture is indeed a wonderful experience. For many of us, these simple rewards are sufficiently gratifying so that we have chosen to spend our scientific lives studying these unusual creatures. In these endeavors many of the strategies and tools as

well as much of the philosophy may be traced to the Delft School, passed on to us by our teachers, Martinus Beijerinck, A. J. Kluyver, and C. B. van Niel, and in turn passed on by us to our students. In this school, the principles of the selective, enrichment culture technique have been developed and diversified; they have been a major force in designing and applying new principles for the capture and isolation of microbes from nature. For me, the “organism approach” has provided rewarding adventures. The organism continually challenges and literally drags the investigator into new areas where unfamiliar tools may be needed. I believe that organismoriented research is an important alternative to problem-oriented research, for new concepts of the future very likely lie in a study of the breadth of microbial life. The physiology, biochemistry, and ecology of the microbe remain the most powerful attractions. Studies based on classical methods as well as modern genetic techniques will result in new insights and concepts. To some readers, this edition of the The Prokaryotes may indicate that the field is now mature, that from here on it is a matter of filling in details. I suspect that this is not the case. Perhaps we have assumed prematurely that we fully understand microbial life. Van Niel pointed out to his students that—after a lifetime of study—it was a very humbling experience to view in the microscope a sample of microbes from nature and recognize only a few. Recent evidence suggests that microbes have been evolving for nearly 4 billion years. Most certainly those microbes now domesticated and kept in captivity in culture collections represent only a minor portion of the species that have evolved in this time span. Sometimes we must remind ourselves that evolution is actively taking place at the present moment. That the eukaryote cell evolved as a chimera of certain prokaryote parts is a generally accepted concept today. Higher as well as lower eukaryotes evolved in contact with prokaryotes, and evidence surrounds us of the complex interactions between eukaryotes and

viii

Foreword

prokaryotes as well as among prokaryotes. We have so far only scratched the surface of these biochemical interrelationships. Perhaps the legume nodule is a pertinent example of nature caught in the act of evolving the “nitrosome,” a unique nitrogen-fixing organelle. Study of prokaryotes is proceeding at such a fast pace that major advances are occurring yearly. The increase of this edition to four volumes documents the exciting pace of discoveries. To prepare a treatise such as The Prokaryotes requires dedicated editors and authors; the task has been enormous. I predict that the scientific community of microbiologists will again show its appreciation through use of these volumes— such that the pages will become “dog-eared” and worn as students seek basic information for the

hunt.These volumes belong in the laboratory, not in the library. I believe that a most effective way to introduce students to microbiology is for them to isolate microbes from nature, i.e., from their habitats in soil, water, clinical specimens, or plants. The Prokaryotes enormously simplifies this process and should encourage the construction of courses that contain a wide spectrum of diverse topics. For the student as well as the advanced investigator these volumes should generate excitement. Happy hunting! Ralph S. Wolfe Department of Microbiology University of Illinois at Urbana-Champaign

Contents

Preface Foreword by Ralph S. Wolfe Contributors

v vii xxix

Volume 1 1. 1.1

Essays in Prokaryotic Biology How We Do, Don’t and Should Look at Bacteria and Bacteriology

3

carl r. woese 1.2

Databases

24

wolfgang ludwig, karl-heinz schleifer and erko stackebrandt 1.3

Defining Taxonomic Ranks

29

erko stackebrandt 1.4

Prokaryote Characterization and Identification

58

hans g. trüper and karl-heinz schleifer 1.5

Principles of Enrichment, Isolation, Cultivation, and Preservation of Prokaryotes

80

jörg overmann 1.6

Prokaryotes and Their Habitats

137

hans g. schlegel and holger w. jannasch 1.7

Morphological and Physiological Diversity

185

stephen h. zinder and martin dworkin 1.8

Cell-Cell Interactions

221

dale kaiser 1.9

Prokaryotic Genomics

246

b. w. wren 1.10

Genomics and Metabolism in Escherichia coli margrethe haugge serres and monica riley

261

x

1.11

Contents

Origin of Life: RNA World versus Autocatalytic Anabolism

275

günter wächtershäuser 1.12

Biotechnology and Applied Microbiology

284

eugene rosenberg 1.13

The Structure and Function of Microbial Communities

299

david a. stahl, meredith hullar and seana davidson

2. 2.1

Symbiotic Associations Cyanobacterial-Plant Symbioses

331

david g. adams, birgitta bergman, s. a. nierzwicki-bauer, a. n. rai and arthur schüßler 2.2

Symbiotic Associations Between Ciliates and Prokaryotes

364

hans-dieter görtz 2.3

Bacteriocyte-Associated Endosymbionts of Insects

403

paul baumann, nancy a. moran and linda baumann 2.4

Symbiotic Associations Between Termites and Prokaryotes

439

andreas brune 2.5

Marine Chemosynthetic Symbioses

475

colleen m. cavanaugh, zoe p. mckiness, irene l.g. newton and frank j. stewart

3. 3.1

Biotechnology and Applied Microbiology Organic Acid and Solvent Production

511

palmer rogers, jiann-shin chen and mary jo zidwick 3.2

Amino Acid Production

756

hidehiko kumagai 3.3

Microbial Exopolysaccharides

766

timothy harrah, bruce panilaitis and david kaplan 3.4

Bacterial Enzymes

777

wim j. quax 3.5

Bacteria in Food and Beverage Production

797

michael p. doyle and jianghong meng 3.6

Bacterial Pharmaceutical Products

812

arnold l. demain and giancarlo lancini 3.7

Biosurfactants

834

eugene rosenberg 3.8

Bioremediation

ronald l. crawford

850

Contents

3.9

Biodeterioration

xi

864

ji-dong gu and ralph mitchell 3.10

Microbial Biofilms

904

dirk de beer and paul stoodley

939

Index

Volume 2 1. 1.1

Ecophysiological and Biochemical Aspects Planktonic Versus Sessile Life of Prokaryotes

3

kevin c. marshall 1.2

Bacterial Adhesion

16

itzhak ofek, nathan sharon and soman n. abraham 1.3

The Phototrophic Way of Life

32

jörg overmann and ferran garcia-pichel 1.4

The Anaerobic Way of Life

86

ruth a. schmitz, rolf daniel, uwe deppenmeier and gerhard gottschalk 1.5

Bacterial Behavior

102

judith armitage 1.6

Prokaryotic Life Cycles

140

martin dworkin 1.7

Life at High Temperatures

167

rainer jaenicke and reinhard sterner 1.8

Life at Low Temperatures

210

siegfried scherer and klaus neuhaus 1.9

Life at High Salt Concentrations

263

aharon oren 1.10

Alkaliphilic Prokaryotes

283

terry ann krulwich 1.11

Syntrophism among Prokaryotes

309

bernhard schink and alfons j.m. stams 1.12

Quorum Sensing

336

bonnie l. bassler and melissa b. miller 1.13

Acetogenic Prokaryotes

harold l. drake, kirsten küsel and carola matthies

354

xii

1.14

Contents

Virulence Strategies of Plant Pathogenic Bacteria

421

barbara n. kunkel and zhongying chen 1.15

The Chemolithotrophic Prokaryotes

441

donovan p. kelly and anne p. wood 1.16

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

457

eberhard bock and michael wagner 1.17

The H2-Metabolizing Prokaryotes

496

edward schwartz and bärbel friedrich 1.18

Hydrocarbon-Oxidizing Bacteria

564

eugene rosenberg 1.19

Cellulose-Decomposing Bacteria and Their Enzyme Systems

578

edward a. bayer, yuval shoham and raphael lamed 1.20

Aerobic Methylotrophic Prokaryotes

618

mary e. lidstrom 1.21

Dissimilatory Fe(III)- and Mn(IV)-Reducing Prokaryotes

635

derek lovley 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

659

ralf rabus, theo a. hansen and friedrich widdel 1.23

The Denitrifying Prokaryotes

769

james p. shapleigh 1.24

Dinitrogen-Fixing Prokaryotes

793

esperanza martinez-romero 1.25

Root and Stem Nodule Bacteria of Legumes

818

michael j. sadowsky and p. h. graham 1.26

Magnetotactic Bacteria

842

stefan spring and dennis a. bazylinski 1.27

Luminous Bacteria

863

paul v. dunlap and kumiko kita-tsukamoto 1.28

Bacterial Toxins

893

vega masignani, mariagrazia pizza and rino rappuoli 1.29

The Metabolic Pathways of Biodegradation

956

lawrence p. wackett 1.30

Haloalkaliphilic Sulfur-Oxidizing Bacteria

969

dimitry yu. sorokin, horia banciu, lesley a. robertson and j. gijs kuenen 1.31

The Colorless Sulfur Bacteria

lesley a. robertson and j. gijs kuenen

985

Contents

1.32

Bacterial Stress Response

xiii

1012

eliora z. ron 1.33

Anaerobic Biodegradation of Hydrocarbons Including Methane

1028

friedrich widdel, antje boetius and ralf rabus 1.34

Physiology and Biochemistry of the Methane-Producing Archaea

1050

reiner hedderich and william b. whitman

1081

Index

Volume 3 A: 1.

Archaea The Archaea: A Personal Overview of the Formative Years

3

ralph s. wolfe 2.

Thermoproteales

10

harald huber, robert huber and karl o. stetter 3.

Sulfolobales

23

harald huber and david prangishvili 4.

Desulfurococcales

52

harald huber and karl o. stetter 5.

The Order Thermococcales

69

costanzo bertoldo and garabed antranikian 6.

The Genus Archaeoglobus

82

patricia hartzell and david w. reed 7.

Thermoplasmatales

101

harald huber and karl o. stetter 8.

The Order Halobacteriales

113

aharon oren 9.

The Methanogenic Bacteria

165

william b. whitman, timothy l. bowen and david r. boone 10.

The Order Methanomicrobiales

208

jean-louis garcia, bernard ollivier and william b. whitman 11.

The Order Methanobacteriales

231

adam s. bonin and david r. boone 12.

The Order Methanosarcinales

melissa m. kendall and david r. boone

244

xiv

13.

Contents

Methanococcales

257

william b. whitman and christian jeanthon 14.

Nanoarchaeota

274

harald huber, michael j. hohn, reinhard rachel and karl o. stetter 15.

Phylogenetic and Ecological Perspectives on Uncultured Crenarchaeota and Korarchaeota scott c. dawson, edward f. delong and norman r. pace

B:

Bacteria

1.

Firmicutes (Gram-Positive Bacteria)

1.1.

Firmicutes with High GC Content of DNA

1.1.1

Introduction to the Taxonomy of Actinobacteria

281

297

erko stackebrandt and peter schumann 1.1.2

The Family Bifidobacteriaceae

322

bruno biavati and paola mattarelli 1.1.3

The Family Propionibacteriaceae: The Genera Friedmanniella, Luteococcus, Microlunatus, Micropruina, Propioniferax, Propionimicrobium and Tessarococcus

383

erko stackebrandt and klaus p. schaal 1.1.4

Family Propionibacteriaceae: The Genus Propionibacterium

400

erko stackebrandt, cecil s. cummins and john l. johnson 1.1.5

The Family Succinivibrionaceae

419

erko stackebrandt and robert b. hespell 1.1.6

1.1.7

The Family Actinomycetaceae: The Genera Actinomyces, Actinobaculum, Arcanobacterium, Varibaculum and Mobiluncus klaus p. schaal, atteyet f. yassin and erko stackebrandt

430

The Family Streptomycetaceae, Part I: Taxonomy

538

peter kämpfer 1.1.8

The Family Streptomycetaceae, Part II: Molecular Biology

605

hildgund schrempf 1.1.9

The Genus Actinoplanes and Related Genera

623

gernot vobis 1.1.10

The Family Actinosynnemataceae

654

david p. labeda 1.1.11

The Families Frankiaceae, Geodermatophilaceae, Acidothermaceae and Sporichthyaceae

philippe normand

669

Contents

1.1.12

The Family Thermomonosporaceae: Actinocorallia, Actinomadura, Spirillospora and Thermomonospora

xv

682

reiner michael kroppenstedt and michael goodfellow 1.1.13

The Family Streptosporangiaceae

725

michael goodfellow and erika teresa quintana 1.1.14

The Family Nocardiopsaceae

754

reiner michael kroppenstedt and lyudmila i. evtushenko 1.1.15

Corynebacterium—Nonmedical

796

wolfgang liebl 1.1.16

The Genus Corynebacterium—Medical

819

alexander von graevenitz and kathryn bernard 1.1.17

The Families Dietziaceae, Gordoniaceae, Nocardiaceae and Tsukamurellaceae

843

michael goodfellow and luis angel maldonado 1.1.18

The Genus Mycobacterium—Nonmedical

889

sybe hartmans, jan a.m. de bont and erko stackebrandt 1.1.19

The Genus Mycobacterium—Medical

919

beatrice saviola and william bishai 1.1.20

Mycobacterium leprae

934

thomas m. shinnick 1.1.21

The Genus Arthrobacter

945

dorothy jones and ronald m. keddie 1.1.22

The Genus Micrococcus

961

miloslav kocur, wesley e. kloos and karl-heinz schleifer 1.1.23

Renibacterium

972

hans-jürgen busse 1.1.24

The Genus Stomatococcus: Rothia mucilaginosa, basonym Stomatococcus mucilaginosus

975

erko stackebrandt 1.1.25

The Family Cellulomonadaceae

983

erko stackebrandt, peter schumann and helmut prauser 1.1.26

The Family Dermatophilaceae

1002

erko stackebrandt 1.1.27

The Genus Brevibacterium

1013

matthew d. collins 1.1.28

The Family Microbacteriaceae

lyudmila i. evtushenko and mariko takeuchi

1020

xvi

1.1.29

Contents

The Genus Nocardioides

1099

jung-hoon yoon and yong-ha park

1115

Index

Volume 4 1.

Firmicutes (Gram-Positive Bacteria)

1.2

Firmicutes with Low GC Content of DNA

1.2.1

The Genera Staphylococcus and Macrococcus

5

friedrich götz, tammy bannerman and karl-heinz schleifer 1.2.2

The Genus Streptococcus—Oral

76

jeremy m. hardie and robert a. whiley 1.2.3

Medically Important Beta-Hemolytic Streptococci

108

p. patrick cleary and qi cheng 1.2.4

Streptococcus pneumoniae

149

elaine tuomanen 1.2.5

The Genus Enterococcus: Taxonomy

163

luc devriese, margo baele and patrick butaye 1.2.6

Enterococcus

175

donald j. leblanc 1.2.7

The Genus Lactococcus

205

michael teuber and arnold geis 1.2.8

The Genera Pediococcus and Tetragenococcus

229

wilhelm h. holzapfel, charles m. a. p. franz, wolfgang ludwig, werner back and leon m. t. dicks 1.2.9

Genera Leuconostoc, Oenococcus and Weissella

267

johanna björkroth and wilhelm h. holzapfel 1.2.10

The Genera Lactobacillus and Carnobacterium

320

walter p. hammes and christian hertel 1.2.11

Listeria monocytogenes and the Genus Listeria

404

nadia khelef, marc lecuit, carmen buchrieser, didier cabanes, olivier dussurget and pascale cossart 1.2.12

The Genus Brochothrix

477

erko stackebrandt and dorothy jones 1.2.13

The Genus Erysipelothrix

erko stackebrandt, annette c. reboli and w. edmund farrar

492

Contents

1.2.14

The Genus Gemella

xvii

511

matthew d. collins 1.2.15

The Genus Kurthia

519

erko stackebrandt, ronald m. keddie and dorothy jones 1.2.16

The Genus Bacillus—Nonmedical

530

ralph a. slepecky and h. ernest hemphill 1.2.17

The Genus Bacillus—Insect Pathogens

563

donald p. stahly, robert e. andrews and allan a. yousten 1.2.18

The Genus Bacillus—Medical

609

w. edmund farrar and annette c. reboli 1.2.19

1.2.20

Genera Related to the Genus Bacillus—Sporolactobacillus, Sporosarcina, Planococcus, Filibacter and Caryophanon dieter claus, dagmar fritze and miloslav kocur

631

An Introduction to the Family Clostridiaceae

654

jürgen wiegel, ralph tanner and fred a. rainey 1.2.21

Neurotoxigenic Clostridia

679

cesare montecucco, ornella rossetto and michel r. popoff 1.2.22

The Enterotoxic Clostridia

698

bruce a. mcclane, francisco a. uzai, mariano e. fernandez miyakawa, david lyerly and tracy wilkins 1.2.23

Clostridium perfringens and Histotoxic Disease

753

julian i. rood 1.2.24

The Genera Desulfitobacterium and Desulfosporosinus: Taxonomy

771

stefan spring and frank rosenzweig 1.2.25

The Genus Desulfotomaculum

787

friedrich widdel 1.2.26

The Anaerobic Gram-Positive Cocci

795

takayuki ezaki, na (michael) li and yoshiaki kawamura 1.2.27

The Order Haloanaerobiales

809

aharon oren 1.2.28

The Genus Eubacterium and Related Genera

823

william g. wade 1.2.29

The Genus Mycoplasma and Related Genera (Class Mollicutes)

836

shmuel razin 1.2.30

The Phytopathogenic Spiroplasmas

jacqueline fletcher, ulrich melcher and astri wayadande

905

xviii

1.3 1.3.1

Contents

Firmicutes with Atypical Cell Walls The Family Heliobacteriaceae

951

michael t. madigan 1.3.2

Pectinatus, Megasphaera and Zymophilus

965

auli haikara and ilkka helander 1.3.3

The Genus Selenomonas

982

robert b. hespell, bruce j. paster and floyd e. dewhirst 1.3.4

The Genus Sporomusa

991

john a. breznak 1.3.5

The Family Lachnospiraceae, Including the Genera Butyrivibrio, Lachnospira and Roseburia

1002

michael cotta and robert forster 1.3.6

The Genus Veillonella

1022

paul kolenbrander 1.3.7

Syntrophomonadaceae

1041

martin sobierj and david r. boone

2. 2.1

Cyanobacteria The Cyanobacteria—Isolation, Purification and Identification

1053

john b. waterbury 2.2

The Cyanobacteria—Ecology, Physiology and Molecular Genetics

1074

yehuda cohen and michael gurevitz 2.3

The Genus Prochlorococcus

1099

anton f. post

1111

Index

Volume 5 3.

Proteobacteria Introduction to the Proteobacteria karel kersters, paul de vos, monique gillis, jean swings,

3

peter van damme and erko stackebrandt

3.1. 3.1.1

Alpha Subclass The Phototrophic Alpha-Proteobacteria

johannes f. imhoff

41

Contents

3.1.2

The Genera Prosthecomicrobium and Ancalomicrobium

xix

65

gary e. oertli, cheryl jenkins, naomi ward, frederick a. rainey, erko stackebrandt and james t. staley 3.1.3

3.1.4

Dimorphic Prosthecate Bacteria: The Genera Caulobacter, Asticcacaulis, Hyphomicrobium, Pedomicrobium, Hyphomonas and Thiodendron jeanne s. poindexter

72

The Genus Agrobacterium

91

ann g. matthysse 3.1.5

The Genus Azospirillum

115

anton hartmann and jose ivo baldani 3.1.6

The Genus Herbaspirillum

141

michael schmid, jose ivo baldani and anton hartmann 3.1.7

The Genus Beijerinckia

151

jan hendrick becking 3.1.8

The Family Acetobacteraceae: The Genera Acetobacter, Acidomonas, Asaia, Gluconacetobacter, Gluconobacter, and Kozakia karel kersters, puspita lisdiyanti, kazuo komagata and

163

jean swings 3.1.9

The Genus Zymomonas

201

hermann sahm, stephanie bringer-meyer and georg a. sprenger 3.1.10

The Manganese-Oxidizing Bacteria

222

kenneth h. nealson 3.1.11

The Genus Paracoccus

232

donovan p. kelly, frederick a. rainey and ann p. wood 3.1.12

The Genus Phenylobacterium

250

jürgen eberspächer and franz lingens 3.1.13

Methylobacterium

257

peter n. green 3.1.14

The Methanotrophs—The Families Methylococcaceae and Methylocystaceae

266

john p. bowman 3.1.15

The Genus Xanthobacter

290

jürgen wiegel 3.1.16

The Genus Brucella

315

edgardo moreno and ignacio moriyón 3.1.17

Introduction to the Rickettsiales and Other Intracellular Prokaryotes

457

david n. fredricks 3.1.18

The Genus Bartonella

michael f. minnick and burt e. anderson

467

xx

Contents

3.1.19

The Order Rickettsiales

493

xue-jie yu and david h. walker 3.1.20

The Genus Coxiella

529

robert a. heinzen and james e. samuel 3.1.21

The Genus Wolbachia

547

markus riegler and scott l. o’neill 3.1.22

Aerobic Phototrophic Proteobacteria

562

vladimir v. yurkov 3.1.23

The Genus Seliberia

585

jean m. schmidt and james r. swafford

3.2. 3.2.1

Beta Subclass The Phototrophic Betaproteobacteria

593

johannes f. imhoff 3.2.2

The Neisseria

602

daniel c. stein 3.2.3

The Genus Bordetella

648

alison weiss 3.2.4

Achromobacter, Alcaligenes and Related Genera

675

hans-jürgen busse and andreas stolz 3.2.5

The Genus Spirillum

701

noel r. krieg 3.2.6

The Genus Aquaspirillum

710

bruno pot, monique gillis and jozef de ley 3.2.7

Comamonas

723

anne willems and paul de vos 3.2.8

The Genera Chromobacterium and Janthinobacterium

737

monique gillis and jozef de ley 3.2.9

The Genera Phyllobacterium and Ochrobactrum

747

jean swings, bart lambert, karel kersters and barry holmes 3.2.10

The Genus Derxia

751

jan hendrick becking 3.2.11

The Genera Leptothrix and Sphaerotilus

758

stefan spring 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

hans-peter koops, ulrike purkhold, andreas pommerening-röser, gabriele timmermann and michael wagner

778

Contents

3.2.13

The Genus Thiobacillus

xxi

812

lesley a. robertson and j. gijs kuenen 3.2.14

The Genera Simonsiella and Alysiella

828

brian p. hedlund and daisy a. kuhn 3.2.15

Eikenella corrodens and Closely Related Bacteria

840

edward j. bottone and paul a. granato 3.2.16

The Genus Burkholderia

848

donald e. woods and pamela a. sokol 3.2.17

The Nitrite-Oxidizing Bacteria

861

aharon abeliovich 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

873

barbara reinhold-hurek and thomas hurek

893

Index

Volume 6 3.

Proteobacteria

3.3.

Gamma Subclass

3.3.1

New Members of the Family Enterobacteriaceae

5

j. michael janda 3.3.2

3.3.3

Phylogenetic Relationships of Bacteria with Special Reference to Endosymbionts and Enteric Species m. pilar francino, scott r. santos and howard ochman

41

The Genus Escherichia

60

rodney a. welch 3.3.4

The Genus Edwardsiella

72

sharon l. abbott and j. michael janda 3.3.5

The Genus Citrobacter

90

diana borenshtein and david b. schauer 3.3.6

The Genus Shigella

99

yves germani and philippe j. sansonetti 3.3.7

The Genus Salmonella

123

craig d. ellermeier and james m. slauch 3.3.8

The Genus Klebsiella

sylvain brisse, francine grimont and patrick a. d. grimont

159

xxii

3.3.9

Contents

The Genus Enterobacter

197

francine grimont and patrick a. d. grimont 3.3.10

The Genus Hafnia

215

megan e. mcbee and david b. schauer 3.3.11

The Genus Serratia

219

francine grimont and patrick a. d. grimont 3.3.12

The Genera Proteus, Providencia, and Morganella

245

jim manos and robert belas 3.3.13

Y. enterocolitica and Y. pseudotuberculosis

270

elisabeth carniel, ingo autenrieth, guy cornelis, hiroshi fukushima, françoise guinet, ralph isberg, jeannette pham, michael prentice, michel simonet, mikael skurnik and georges wauters 3.3.14

Yersinia pestis and Bubonic Plague

399

robert brubaker 3.3.15

Erwinia and Related Genera

443

clarence i. kado 3.3.16

The Genera Photorhabdus and Xenorhabdus

451

noel boemare and raymond akhurst 3.3.17

The Family Vibrionaceae

495

j. j. farmer, iii 3.3.18

The Genera Vibrio and Photobacterium

508

j. j. farmer, iii and f. w. hickman-brenner 3.3.19

The Genera Aeromonas and Plesiomonas

564

j. j. farmer, iii, m. j. arduino and f. w. hickman-brenner 3.3.20

The Genus Alteromonas and Related Proteobacteria

597

valery v. mikhailov, lyudmila a. romanenko and elena p. ivanova 3.3.21

Nonmedical: Pseudomonas

646

edward r. b. moore, brian j. tindall, vitor a. p. martins dos santos, dietmar h. pieper, juan-luis ramos and norberto j. palleroni 3.3.22

Pseudomonas aeruginosa

704

timothy l. yahr and matthew r. parsek 3.3.23

Phytopathogenic Pseudomonads and Related Plant-Associated Pseudomonads milton n. schroth, donald c. hildebrand and

714

nickolas panopoulos 3.3.24

Xylophilus

anne willems and monique gillis

741

Contents

3.3.25

The Genus Acinetobacter

xxiii

746

kevin towner 3.3.26

The Family Azotobacteraceae

759

jan hendrick becking 3.3.27

The Genera Beggiatoa and Thioploca

784

andreas teske and douglas c. nelson 3.3.28

The Family Halomonadaceae

811

david r. arahal and antonio ventosa 3.3.29

The Genus Deleya

836

karel kersters 3.3.30

The Genus Frateuria

844

jean swings 3.3.31

The Chromatiaceae

846

johannes f. imhoff 3.3.32

The Family Ectothiorhodospiraceae

874

johannes f. imhoff 3.3.33

Oceanospirillum and Related Genera

887

josé m. gonzález and william b. whitman 3.3.34

Serpens flexibilis: An Unusually Flexible Bacterium

916

robert b. hespell 3.3.35

The Genus Psychrobacter

920

john p. bowman 3.3.36

The Genus Leucothrix

931

thomas d. brock 3.3.37

The Genus Lysobacter

939

hans reichenbach 3.3.38

The Genus Moraxella

958

john p. hays 3.3.39

Legionella Species and Legionnaire’s Disease

988

paul h. edelstein and nicholas p. cianciotto 3.3.40

The Genus Haemophilus

1034

doran l. fink and joseph w. st. geme, iii 3.3.41

The Genus Pasteurella

1062

henrik christensen and magne bisgaard 3.3.42

The Genus Cardiobacterium

sydney m. harvey and james r. greenwood

1091

xxiv

3.3.43

Contents

The Genus Actinobacillus

1094

janet i. macinnes and edward t. lally 3.3.44

The Genus Francisella

1119

francis nano and karen elkins 3.3.45

Ecophysiology of the Genus Shewanella

1133

kenneth h. nealson and james scott 3.3.46

The Genus Nevskia

1152

heribert cypionka, hans-dietrich babenzien, frank oliver glöckner and rudolf amann 3.3.47

The Genus Thiomargarita

1156

heide n. schulz

1165

Index

Volume 7 3.

Proteobacteria

3.4

Delta Subclass

3.4.1

The Genus Pelobacter

5

bernhard schink 3.4.2

The Genus Bdellovibrio

12

edouard jurkevitch 3.4.3

The Myxobacteria

31

lawrence j. shimkets, martin dworkin and hans reichenbach

3.5. 3.5.1

Epsilon Subclass The Genus Campylobacter

119

trudy m. wassenaar and diane g. newell 3.5.2

The Genus Helicobacter

139

jay v. solnick, jani l. o’rourke, peter van damme and adrian lee 3.5.3

The Genus Wolinella

178

jörg simon, roland gross, oliver klimmek and achim kröger

4. 4.1

Spirochetes Free-Living Saccharolytic Spirochetes: The Genus Spirochaeta

susan leschine, bruce j. paster and ercole canale-parola

195

Contents

4.2

The Genus Treponema

xxv

211

steven j. norris, bruce j. paster, annette moter and ulf b. göbel 4.3

The Genus Borrelia

235

melissa j. caimano 4.4

The Genus Leptospira

294

ben adler and solly faine 4.5

Termite Gut Spirochetes

318

john a. breznak and jared r. leadbetter 4.6

The Genus Brachyspira

330

thaddeus b. stanton

5. 5.1

Chlorobiaceae The Family Chlorobiaceae

359

jörg overmann

6. 6.1

Bacteroides and Cytophaga Group The Medically Important Bacteroides spp. in Health and Disease

381

c. jeffrey smith, edson r. rocha and bruce j. paster 6.2

The Genus Porphyromonas

428

frank c. gibson and caroline attardo genco 6.3

An Introduction to the Family Flavobacteriaceae

455

jean-françois bernardet and yasuyoshi nakagawa 6.4

The Genus Flavobacterium

481

jean-françois bernardet and john p. bowman 6.5

The Genera Bergeyella and Weeksella

532

celia j. hugo, brita bruun and piet j. jooste 6.6

The Genera Flavobacterium, Sphingobacterium and Weeksella

539

barry holmes 6.7

The Order Cytophagales

549

hans reichenbach 6.8

The Genus Saprospira

591

hans reichenbach 6.9

The Genus Haliscomenobacter

602

eppe gerke mulder and maria h. deinema 6.10

Sphingomonas and Related Genera

david l. balkwill, j. k. fredrickson and m. f. romine

605

xxvi

6.11

Contents

The Genera Empedobacter and Myroides

630

celia j. hugo, brita bruun and piet j. jooste 6.12

The Genera Chryseobacterium and Elizabethkingia

638

jean-françois bernardet, celia j. hugo and brita bruun 6.13

6.14

6.15

The Marine Clade of the Family Flavobacteriaceae: The Genera Aequorivita, Arenibacter, Cellulophaga, Croceibacter, Formosa, Gelidibacter, Gillisia, Maribacter, Mesonia, Muricauda, Polaribacter, Psychroflexus, Psychroserpens, Robiginitalea, Salegentibacter, Tenacibaculum, Ulvibacter, Vitellibacter and Zobellia john p. bowman

677

Capnophilic Bird Pathogens in the Family Flavobacteriaceae: Riemerella, Ornithobacterium and Coenonia peter van damme, h. m. hafez and k. h. hinz

695

The Genus Capnocytophaga

709

e. r. leadbetter 6.16

The Genera Rhodothermus, Thermonema, Hymenobacter and Salinibacter

712

aharon oren

7. 7.1

Chlamydia The Genus Chlamydia—Medical

741

murat v. kalayoglu and gerald i. byrne

8.

Planctomyces and Related Bacteria

8.1

The Order Planctomycetales, Including the Genera Planctomyces, Pirellula, Gemmata and Isosphaera and the Candidatus Genera Brocadia, Kuenenia and Scalindua naomi ward, james t. staley, john a. fuerst, stephen giovannoni,

757

heinz schlesner and erko stackebrandt

9. 9.1

Thermus The Genus Thermus and Relatives

797

milton s. da costa, frederick a. rainey and m. fernanda nobre

10. 10.1

Chloroflexaceae and Related Bacteria The Family Chloroflexaceae

815

satoshi hanada and beverly k. pierson 10.2

The Genus Thermoleophilum

843

jerome j. perry 10.3

The Genus Thermomicrobium

jerome j. perry

849

Contents

10.4

The Genus Herpetosiphon

xxvii

854

natuschka lee and hans reichenbach

11.

Verrucomicrobium

11.1

The Phylum Verrucomicrobia: A Phylogenetically Heterogeneous Bacterial Group heinz schlesner, cheryl jenkins and james t. staley

12. 12.1

881

Thermotogales Thermotogales

899

robert huber and michael hannig

13. 13.1

Aquificales Aquificales

925

robert huber and wolfgang eder

14. 14.1

Phylogenetically Unaffiliated Bacteria Morphologically Conspicuous Sulfur-Oxidizing Eubacteria

941

jan w. m. la rivière and karin schmidt 14.2

The Genus Propionigenium

955

bernhard schink 14.3

The Genus Zoogloea

960

patrick r. dugan, daphne l. stoner and harvey m. pickrum 14.4

Large Symbiotic Spirochetes: Clevelandina, Cristispira, Diplocalyx, Hollandina and Pillotina

971

lynn margulis and gregory hinkle 14.5

Streptobacillus moniliformis

983

james r. greenwood and sydney m. harvey 14.6

The Genus Toxothrix

986

peter hirsch 14.7

The Genus Gallionella

990

hans h. hanert 14.8

The Genera Caulococcus and Kusnezovia

996

jean m. schmidt and georgi a. zavarzin 14.9

The Genus Brachyarcus

peter hirsch

998

xxviii

14.10

Contents

The Genus Pelosigma

1001

peter hirsch 14.11

14.12

The Genus Siderocapsa (and Other Iron- and Maganese-Oxidizing Eubacteria) hans h. hanert

1005

The Genus Fusobacterium

1016

tor hofstad 14.13

Prokaryotic Symbionts of Amoebae and Flagellates

1028

kwang w. jeon Index

1039

Contributors

Sharon L. Abbott Microbial Diseases Laboratory Berkeley, CA 94704 USA Aharon Abeliovich Department of Biotechnology Engineering Institute for Applied Biological Research Environmental Biotechnology Institute Ben Gurion University 84105 Beer-Sheva Israel Soman N. Abraham Director of Graduate Studies in Pathology Departments of Pathology, Molecular Genetics and Microbiology, and Immunology Duke University Medical Center Durham, NC 27710 USA David G. Adams School of Biochemistry and Microbiology University of Leeds Leeds LS2 9JT UK Ben Adler Monash University Faculty of Medicine, Nursing and Health Sciences Department of Microbiology Clayton Campus Victoria, 3800 Australia Raymond Akhurst CSIRO Entomology Black Mountain ACT 2601 Canberra Australia Rudolf Amann Max Planck Institute for Marine Microbiology D-28359 Bremen Germany

Burt E. Anderson Department of Medical Microbiology and Immunology College of Medicine University of South Florida Tampa, FL 33612 USA Robert E. Andrews Department of Microbiology University of Iowa Iowa City, IA 52242 USA Garabed Antranikian Technical University Hamburg-Harburg Institute of Technical Microbiology D-21073 Hamburg Germany David R. Arahal Colección Española de Cultivos Tipo (CECT) Universidad de Valencia Edificio de Investigación 46100 Burjassot (Valencia) Spain M. J. Arduino Center for Infectious Diseases Centers for Disease Control Atlanta, GA 30333 USA Judith Armitage Department of Biochemistry Microbiology Unit University of Oxford OX1 3QU Oxford UK Ingo Autenrieth Institut für Medizinische Mikrobiologie Universitatsklinikum Tuebingen D-72076 Tuebingen Germany

xxx

Contributors

Hans-Dietrich Babenzien Leibniz-Institut für Gewässerökologie und Binnenfischereiim Forschungsverbund Berlin 12587 Berlin Germany

Paul Baumann Department of Microbiology University of California, Davis Davis, CA 95616-5224 USA

Werner Back Lehrstuhl für Technologie der Brauerei I Technische Universität München D-85354 Freising-Weihenstephan Germany

Edward A. Bayer Department of Biological Chemistry Weizmann Institute of Science Rehovot 76100 Israel

Margo Baele Department of Pathology Bacteriology and Poultry Diseases Faculty of Veterinary Medicine Ghent University B-9820 Merelbeke Belgium

Dennis A. Bazylinski Department of Microbiology, Immunology and Preventive Medicine Iowa State University Ames, IA 55001 USA

Jose Ivo Baldani EMBRAPA-Agrobiology Centro Nacional de Pesquisa de Agrobiologia Seropedica, 23851-970 CP 74505 Rio de Janeiro Brazil

Jan Hendrick Becking Stichting ITAL Research Institute of the Ministry of Agriculture and Fisheries 6700 AA Wageningen The Netherlands

David L. Balkwill Department of Biomedical Sciences College of Medicine Florida State University Tallahassee, FL 32306-4300 USA Horia Banciu Department of Biotechnology Delft University of Technology 2628 BC Delft Tammy Bannerman School of Allied Medical Professions Division of Medical Technology The Ohio State University Columbus, OH 43210 USA Bonnie L. Bassler Department of Molecular Biology Princeton University Princeton, NJ 08544-1014 USA Linda Baumann School of Nursing Clinical Science Center University of Wisconsin Madison, WI 53792-2455 USA

Robert Belas The University of Maryland Biotechnology Institute Center of Marine Biotechnology Baltimore, MD 21202 USA Birgitta Bergman Department of Botany Stockholm University SE-106 91 Stockholm Sweden Kathryn Bernard Special Bacteriology Section National Microbiology Laboratory Health Canada Winnipeg R3E 3R2 Canada Jean-François Bernardet Unité de Virologie et Immunologie Moléculaires Institut National de la Recherche Agronomique (INRA) Domaine de Vilvert 78352 Jouy-en-Josas cedex France

Contributors

Costanzo Bertoldo Technical University Hamburg-Harburg Institute of Technical Microbiology D-21073 Hamburg Germany Bruno Biavati Istituto di Microbiologia Agraria 40126 Bologna Italy Magne Bisgaard Department of Veterinary Microbiology Royal Veterinary and Agricultural University 1870 Frederiksberg C Denmark William Bishai Departments of Molecular Microbiology and Immunology, International Health, and Medicine Center for Tuberculosis Research Johns Hopkins School of Hygiene and Public Health Baltimore, MD 21205-2105 USA Johanna Björkroth Department of Food and Environmental Hygiene Faculty of Veterinary Medicine University of Helsinki FIN-00014 Helsinki Finland Eberhard Bock Institute of General Botany Department of Microbiology University of Hamburg D-22609 Hamburg Germany Noel Boemare Ecologie Microbienne des Insectes et Interactions Hôte-Pathogène UMR EMIP INRA-UMII IFR56 Biologie cellulaire et Porcessus infectieux Université Montpellier II 34095 Montpellier France Antje Boetius Max-Planck-Institut für Marine Mikrobiologie D-28359 Bremen Germany Adam S. Bonin Portland State University Portland OR 97207 USA

xxxi

David R. Boone Department of Biology Environmental Science and Engineering Oregon Graduate Institute of Science and Technology Portland State University Portland, OR 97207-0751 USA Diana Borenshtein Massachusetts Institute of Technology Cambridge, MA 02139-4307 USA Edward J. Bottone Division of Infectious Diseases The Mount Sinai Hospital One Gustave L. Levy Place New York, NY 10029 USA Timothy L. Bowen Department of Microbiology University of Georgia Athens, GA 30602 USA John P. Bowman Australian Food Safety Centre for Excellence School of Agricultural Science Hobart, Tasmania, 7001 Australia John A. Breznak Department of Microbiology and Molecular Genetics Michigan State University East Lansing, MI 48824-1101 USA Stephanie Bringer-Meyer Institut Biotechnologie Forschungszentrum Jülich D-52425 Jülich Germany Sylvain Brisse Unité Biodiversité des Bactéries Pathogènes Emergentes U 389 INSERM Institut Pasteur 75724 Paris France Thomas D. Brock Department of Bacteriology University of Wisconsin-Madison Madison, WI 53706 USA

xxxii

Contributors

Robert Brubaker Department of Microbiology Michigan State University East Lansing, MI 48824 USA

Ercole Canale-Parola Department of Microbiology University of Massachusetts Amherst, MA 01003 USA

Andreas Brune Max Planck Institute for Terrestrial Microbiology Marburg Germany

Elisabeth Carniel Laboratoire des Yersinia Institut Pasteur 75724 Paris France

Brita Bruun Department of Clinical Microbiology Hillerød Hospital DK 3400 Hillerød Denmark Carmen Buchrieser Laboratoire de Génomique des Microorganismes Pathogènes Institut Pasteur 75724 Paris France Hans-Jürgen Busse Institut für Bakteriology, Mykologie, und Hygiene Veterinärmedizinische Universität Wien A-1210 Vienna Austria Patrick Butaye CODA-CERVA-VAR 1180 Brussels Belgium Gerald I. Byrne Department of Medical Microbiology and Immunology University of Wisconsin—Madison Madison, WI 53706 USA Didier Cabanes Department of Immunology and Biology of Infection Molecular Microbiology Group Institute for Molecular and Cellular Biology 4150-180 Porto Portugal Melissa Caimano Center for Microbial Pathogenesis and Department of Pathology and Department of Genetics and Development University of Connecticut Health Center Farmington, CT 06030-3205 USA

Colleen M. Cavanaugh Bio Labs Harvard University Cambridge, MA 02138 USA Jiann-Shin Chen Department of Biochemistry Virginia Polytechnic Institute and State University—Virginia Tech Blacksburg, VA 24061-0308 USA Zhongying Chen Department of Biology University of North Carolina Chapel Hill, NC 27514 USA Qi Cheng University of Western Sydney Penrith South NSW 1797 Australia Henrik Christensen Department of Veterinary Microbiology Royal Veterinary and Agricultural University Denmark Nicholas P. Cianciotto Department of Microbiology and Immunology Northwestern University School of Medicine Chicago, IL USA Dieter Claus Deutsche Sammlung von Mikroorganismen D-3300 Braunschweig-Stockheim Germany P. Patrick Cleary Department of Microbiology University of Minnesota Medical School Minneapolis, MN 55455 USA

Contributors

xxxiii

Yehuda Cohen Department of Molecular and Microbial Ecology Institute of Life Science Hebrew University of Jerusalem 91904 Jerusalem Israel

Milton S. da Costa M. Fernanda Nobre Centro de Neurociências e Biologia Celular Departamento de Zoologia Universidade de Coimbra 3004-517 Coimbra Portugal

Matthew D. Collins Institute of Food Research Reading Lab, Early Gate UK

Rolf Daniel Department of General Microbiology Institute of Microbiology and Genetics 37077 Göttingen Germany

Guy Cornelis Microbial Pathogenesis Unit Université Catholique de Louvain and Christian de Duve Institute of Cellular Pathology B1200 Brussels Belgium

Seana Davidson University of Washington Civil and Environmental Engineering Seattle, WA 98195-2700 USA

Pascale Cossart Unité des Interactions Bactéries-Cellules INSERM U604 Institut Pasteur 75724 Paris France Michael Cotta USDA-ARS North Regional Research Center Peoria, IL 61604-3902 USA Ronald L. Crawford Food Research Center University of Idaho Moscow, ID 83844-1052 USA Cecil S. Cummins Department of Anaerobic Microbiology Virginia Polytechnic Institute and State University Blacksburg, VA 24061 USA Heribert Cypionka Institut für Chemie und Biologie des Meeres Fakultät 5, Mathematik und Naturwissenschaften Universität Oldenburg D-26111 Oldenburg Germany

Scott C. Dawson Department of Molecular and Cellular Biology University of California-Berkeley Berkeley, CA 94720 USA Dirk de Beer Max-Planck-Institute for Marine Microbiology D-28359 Bremen Germany Jan A.M. de Bont Department of Food Science Agricultural University 6700 EV Wageningen The Netherlands Maria H. Deinema Laboratory of Microbiology Agricultural University 6703 CT Wageningen The Netherlands Jozef de Ley Laboratorium voor Microbiologie en Microbiële Genetica Rijksuniversiteit Ghent B-9000 Ghent Belgium Edward F. DeLong Science Chair Monterey Bay Aquarium Research Institute Moss Landing, CA 95039 USA

xxxiv

Contributors

Arnold L. Demain Department of Biology Massachusetts Institute of Technology Cambridge, MA 02139 USA Uwe Deppenmeier Department of Biological Sciences University of Wisconsin Milwaukee, WI 53202 USA Paul de Vos Department of Biochemistry, Physiology and Microbiology Universiteit Gent B-9000 Gent Belgium Luc Devriese Faculty of Veterinary Medicine B982 Merelbeke Belgium Floyd E. Dewhirst Forsyth Dental Center 140 Fenway Boston, MA 02115 USA Leon M. T. Dicks Department of Microbiology University of Stellenbosch ZA-7600 Stellenbosch South Africa Michael P. Doyle College of Agricultural and Environmental Sciences Center for Food Safety and Quality Enhancement University of Georgia Griffin, GA 30223-1797 USA Harold L. Drake Department of Ecological Microbiology BITOEK, University of Bayreuth D-95440 Bayreuth Germany Patrick R. Dugan Idaho National Engineering Laboratory EG & G Idaho Idaho Falls, ID 83415 USA

Paul V. Dunlap Department of Molecular Cellular and Developmental Biology University of Michigan Ann Arbor, MI 48109-1048 USA Olivier Dussurget Unité des Interactions Bactéries-Cellules INSERM U604 Institut Pasteur 75724 Paris France Martin Dworkin University of Minnesota Medical School Department of Microbiology Minneapolis, MN 55455 USA Jürgen Eberspächer Institut fur Mikrobiologie Universitat Hohenheim D-7000 Stuttgart 70 Germany Paul H. Edelstein Department of Pathology and Laboratory Medicine University of Pennsylvania Medical Center Philadelphia, PA 19104-4283 USA Wolfgang Eder Lehrstuhl für Mikrobiologie Universität Regensburg 93053 Regensburg Germany Karen Elkins CBER/FDA Rockville, MD 20852 USA Craig D. Ellermeier Department of Microbiology University of Illinois Urbana, IL 61801 and Department of Molecular and Cellular Biology Harvard University Cambridge, MA 02138 USA

Contributors

Lyudmila I. Evtushenko All-Russian Collection of Microorganisms Institute of Biochemistry and Physiology of the Russian, Academy of Sciences Puschino Moscow Region, 142290 Russia Takayuki Ezaki Bacterial Department Gifu University Medical School 40 Tsukasa Machi Gifu City Japan Solly Faine Monash University Faculty of Medicine, Nursing and Health Sciences Department of Microbiology Clayton Campus Victoria, 3800 Australia J. J. Farmer, III Center for Infectious Diseases Centers for Disease Control Atlanta, GA 30333 USA W. Edmund Farrar Department of Medicine Medical University of South Carolina Charleston, SC 29425 USA Mariano E. Fernandez Miyakawa California Animal Health and Food Safety Laboratory University of California, Davis San Bernardino, CA 92408 USA

Robert Forster Bio-Products and Bio-Processes Program Agriculture and Agri-Food Canada Lethbridge Research Centre Lethbridge T1J 4B1 Canada M. Pilar Francino Evolutionary Genomics Department DOE Joint Genome Institute Walnut Creek, CA 94598 USA Charles M. A. P. Franz Institute of Hygiene and Toxicology BFEL D-76131 Karlsruhe Germany David N. Fredricks VA Palo Alto Healthcare System Palo Alto, CA 94304 USA J. K. Fredrickson Pacific Northwest National Laboratory Richland, Washington 99352 USA Bärbel Friedrich Institut für Biologie/Mikrobiologie Homboldt-Universität zu Berlin Chaussesstr. 117 D-10115 Berlin Germany Dagmar Fritze Deutsche Sammlung von Mikroorganismen D-3300 Braunschweig-Stockheim Germany

Doran L. Fink Edward Mallinckrodt Department of Pediatrics and Department of Molecular Microbiology Washington University School of Medicine St. Louis, Missouri 63110 USA

John A. Fuerst Department of Microbiology and Parasitology University of Queensland Brisbane Queensland 4072 Australia

Jacqueline Fletcher Department of Entomology and Plant Pathology Oklahoma State University Stillwater, OK USA

Hiroshi Fukushima Public Health Institute of Shimane Prefecture 582-1 Nishihamasada, Matsue Shimane 690-0122 Japan

xxxv

xxxvi

Contributors

Jean-Louis Garcia Laboratoire ORSTOM de Microbiologie des Anaérobies Université de Provence CESB-ESIL 13288 Marseille France

Ulf B. Göbel Institut für Mikrobiologie und Hygiene Universitaetsklinikum Chariteacute Humboldt-Universitaet zu Berlin D-10117 Berlin Germany

Ferran Garcia-Pichel Associate Professor Arizona State University Tempe, AZ 85281 USA

José M. González Department de Microbiologia y Biologia Celular Facultad de Farmacia Universidad de La Laguna 38071 La Laguna, Tenerife SPAIN

Arnold Geis Institut für Mikrobiologie Bundesanstalt für Milchforschung D-24121 Kiel Germany Caroline Attardo Genco Department of Medicine Section of Infectious Diseases and Department of Microbiology Boston University School of Medicine Boston, MA 02118 USA Yves Germani Institut Pasteur Unité Pathogénie Microbienne Moléculaire and Réseau International des Instituts Pasteur Paris 15 France Frank C. Gibson Department of Medicine Section of Infectious Diseases and ‘ Department of Microbiology Boston University School of Medicine Boston, MA 02118 USA Monique Gillis Laboratorium voor Mikrobiologie Universiteit Gent B-9000 Gent Belgium Stephen Giovannoni Department of Microbiology Oregon State University Corvallis, OR 97331 USA Frank Oliver Glöckner Max-Planck-Institut für Marine Mikrobiologie D-28359 Bremen Germany

Michael Goodfellow School of Biology Universtiy of Newcastle Newcastle upon Tyre NE1 7RU UK Friedrich Götz Facultät für Biologie Institut für Microbielle Genetik Universität Tübingen D-72076 Tübingen Germany Hans-Dieter Görtz Department of Zoology Biologisches Institut Universität Stuttgart D-70569 Stuttgart Germany Gerhard Gottschalk Institut für Mikrobiologie und Genetik Georg-August-Universität Göttingen D-37077 Göttingen Germany P. H. Graham Department of Soil, Water, and Climate St. Paul, MN 55108 USA Paul A. Granato Department of Microbiology and Immunology State University of New York Upstate Medical University Syracus, NY 13210 USA Peter N. Green NCIMB Ltd AB24 3RY Aberdeen UK

Contributors

James R. Greenwood Bio-Diagnostics Laboratories Torrance, CA 90503 USA Francine Grimont Unite 199 INSERM Institut Pasteur 75724 Paris France Patrick A. D. Grimont Institut Pasteur 75724 Paris France Roland Gross Institut für Mikrobiologie Johann Wolfgang Goethe-Universität Frankfurt am Main Germany Ji-Dong Gu Laboratory of Environmental Toxicology Department of Ecology & Biodiversity and The Swire Institute of Marine Science University of Hong Kong Hong Kong SAR P.R. China and Environmental and Molecular Microbiology South China Sea Institute of Oceanography Chinese Academy of Sciences Guangzhou 510301 P.R. China Françoise Guinet Laboratoire des Yersinia Institut Pasteur 75724 Paris France Michael Gurevitz Department of Botany Life Sciences Institute Tel Aviv University Ramat Aviv 69978 Israel H. M. Hafez Institute of Poultry Diseases Free University Berlin Berlin German Auli Haikara VTT Biotechnology Tietotie 2, Espoo Finland

Walter P. Hammes Institute of Food Technology Universität Hohenheim D-70599 Stuttgart Germany Satoshi Hanada Research Institute of Biological Resources National Institute of Advanced Industrial Science and Technology (AIST) Tsukuba 305-8566 Japan Hans H. Hanert Institut für Mikrobiologie Technische Univeristät Braunschweig D-3300 Braunschweig Germany Michael Hannig Lehrstuhl für Mikrobiologie Universität Regensburg D-93053 Regensburg Germany Theo A. Hansen Microbial Physiology (MICFYS) Groningen University Rijksuniversiteit Groningen NL-9700 AB Groningen The Netherlands Jeremy M. Hardie Department of Oral Microbiology School of Medicine & Dentistry London E1 2AD UK Timothy Harrah Bioengineering Center Tufts University Medford, MA 02155 USA Anton Hartmann GSF-National Research Center for Environment and Health Institute of Soil Ecology Rhizosphere Biology Division D-85764 Neuherberg/Muenchen Germany Sybe Hartmans Department of Food Science Agricultural University Wageningen 6700 EV Wageningen The Netherlands

xxxvii

xxxviii

Contributors

Patricia Hartzell Department of Microbiology, Molecular Biology, and Biochemistry University of Idaho Moscow, ID 83844-3052 USA

F. W. Hickman-Brenner Center for Infectious Diseases Centers for Disease Control Atlanta, GA 30333 USA

Sydney M. Harvey Nichols Institute Reference Laboratories 32961 Calle Perfecto San Juan Capistrano, CA 92675 USA

Donald C. Hildebrand Department of Plant Pathology University of California-Berkeley Berkeley, CA 94720 USA

John P. Hays Department of Medical Microbiology and Infectious Diseases Erasmus MC 3015 GD Rotterdam The Netherlands

Gregory Hinkle Department of Botany University of Massachusetts Amherst, MA 01003 USA

Reiner Hedderich Max Planck Institute für Terrestriche Mikrobiologie D-35043 Marburg Germany

K. H. Hinz Clinic for Poultry School of Veterinary Medicine D-30559 Hannover Germany

Brian P. Hedlund Department of Biological Sciences University of Nevada, Las Vegas Las Vegas, NV 89154-4004 USA Robert A. Heinzen Department of Molecular Biology University of Wyoming Laramie, WY 82071-3944 USA Ilkka Helander VTT Biotechnology Tietotie 2, Espoo Finland H. Ernest Hemphill Department of Biology Syracuse University Syracuse, NY 13244 USA Christian Hertel Institute of Food Technology Universität Hohenheim D-70599 Stuttgart Germany Robert B. Hespell Northern Regional Research Center, ARS US Department of Agriculture Peoria, IL 61604 USA

Peter Hirsch Institut für Allgemeine Mikrobiologie Universität Kiel D-2300 Kiel Germany Tor Hofstad Department of Microbiology and Immunology University of Bergen N-5021 Bergen Norway Michael J. Hohn Lehrstuhl für Mikrobiologie Universität Regensburg D-93053 Regensburg Germany Barry Holmes Central Public Health Laboratory National Collection of Type Cultures London NW9 5HT UK Wilhelm H. Holzapfel Federal Research Centre of Nutrition Institute of Hygiene and Toxicology D-76131 Karlsruhe Germany

Contributors

Harald Huber Lehrstuhl für Mikrobiologie Universität Regensburg D-93053 Regensburg Germany

Robert Huber Lehrstuhl für Mikrobiologie Universität Regensburg D-93053 Regensburg Germany

Celia J. Hugo Department of Microbial, Biochemical and Food Biotechnology University of the Free State Bloemfontein South Africa

Meredith Hullar University of Washington Seattle, WA USA Thomas Hurek Laboratory of General Microbiology University Bremen 28334 Bremen Germany Johannes F. Imhoff Marine Mikrobiologie Institut für Meereskunde an der Universität Kiel D-24105 Kiel Germany Ralph Isberg Department of Molecular Biology and Microbiology Tufts University School of Medicine Boston, MA 02111 USA

Elena P. Ivanova Senior Researcher in Biology Laboratory of Microbiology Pacific Institute of Bioorganic Chemistry of the Far-Eastern Branch of the Russian Academy of Sciences 690022 Vladivostok Russia

xxxix

Rainer Jaenicke 6885824 Schwalbach a. Ts. Germany and Institut für Biophysik und Physikalische Biochemie Universität Regensburg Regensburg Germany and School of Crystallography Birbeck College University of London London, UK

J. Michael Janda Microbial Diseases Laboratory Division of Communicable Disease Control California Department of Health Services Berkeley, CA 94704-1011 USA

Holger W. Jannasch Woods Hole Oceanographic Institution Woods Hole, MA 02543 USA

Christian Jeanthon UMR CNRS 6539–LEMAR Institut Universitaire Europeen de la Mer Technopole Brest Iroise 29280 Plouzane France

Cheryl Jenkins Department of Microbiology University of Washington Seattle, WA 98195 USA

John L. Johnson Department of Anaerobic Microbiology Virginia Polytechnic Institute and State University Blacksburg, VA 24061 USA

Dorothy Jones Department of Microbiology University of Leicester, School of Medicine Lancaster LE1 9HN UK

xl

Contributors

Piet J. Jooste Department of Biotechnology and Food Technology Tshwane University of Technology Pretoria 0001 South Africa Edouard Jurkevitch Department of Plant Pathology and Microbiology Faculty of Agriculture Food & Environmental Quality Services The Hebrew University 76100 Rehovot Israel Clarence I. Kado Department of Plant Pathology University of California, Davis Davis, CA 95616-5224 USA Dale Kaiser Department of Biochemistry Stanford University School of Medicine Stanford, CA 94305-5329 USA Murat V. Kalayoglu Department of Medical Microbiology and Immunology University of Wisconsin—Madison Madison, WI 53706 USA Peter Kämpfer Institut für Angewandte Mikrobiologie Justus Liebig-Universität D-35392 Gießen Germany David Kaplan Department of Chemcial and Biological Engineering Tufts University Medford, MA 02115 USA Yoshiaki Kawamura Department of Microbiology Regeneration and Advanced Medical Science Gifu University Graduate School of Medicine Gifu 501-1194 Japan

Ronald M. Keddie Craigdhu Fortrose Ross-shire IV 10 8SS UK Donovan P. Kelly University of Warwick Department of Biological Sciences CV4 7AL Coventry UK Melissa M. Kendall Department of Biology Portland State University Portland, OR 97207-0751 USA Karel Kersters Laboratorium voor Mikrobiologie Department of Biochemistry Physiology and Microbiology Universiteit Gent B-9000 Gent Belgium Nadia Khelef Unité des Interactions Bactéries-Cellules INSERM U604 Institut Pasteur 75724 Paris France Kumiko Kita-Tsukamoto Ocean Research Institute University of Tokyo Tokyo 164 Japan Oliver Klimmek Johann Wolfgang Goethe-Universität Frankfurt Institut für Mikrobiologie D-60439 Frankfurt Germany Wesley E. Kloos Department of Genetics North Carolina State University Raleigh, NC 27695-7614 USA Miloslav Kocur Czechoslovak Collection of Microorganisms J.E. Purkyneˇ University 662 43 Brno Czechoslovakia

Contributors

Paul Kolenbrander National Institute of Dental Research National Institute of Health Bethesda, MD 20892-4350 USA Kazuo Komagata Laboratory of General and Applied Microbiology Department of Applied Biology and Chemistry Faculty of Applied Bioscience Tokyo University of Agriculture Tokyo, Japan Hans-Peter Koops Institut für Allgemeine Botanik Abteilung Mikrobiologie Universität Hamburg D-22069 Hamburg Germany Noel R. Krieg Department of Biology Virginia Polytechnic Institute Blacksburg, VA 24061-0406 USA Achim Kröger Institut für Mikrobiologie Biozentrum Niederursel D-60439 Frankfurt/Main Germany

Hidehiko Kumagai Division of Applied Sciences Graduate School of Agriculture Kyoto University Kitashirakawa 606 8502 Kyoto Japan Barbara N. Kunkel Department of Biology Washington University St. Louis, MO 63130 USA Kirsten Küsel Department of Ecological Microbiology BITOEK, University of Bayreuth D-95440 Bayreuth Germany David P. Labeda Microbial Genomics and Bioprocessing Research Unit National Center for Agricultural Utilization Research Agricultural Research Service U.S. Department of Agriculture Peoria, IL 61604 USA Edward T. Lally Leon Levy Research Center for Oral Biology University of Pennsylvania Philadelphia, Pennsylvania, 19104-6002 USA

Reiner Michael Kroppenstedt Deutsche Sammlung von Mikroorganismen und Zellkulturen D-3300 Braunschweig Germany

Bart Lambert Plant Genetic Systems N.V. J. Plateaustraat 22 B-9000 Ghent Belgium

Terry Ann Krulwich Department of Biochemistry Mount Sinai School of Medicine New York, NY 10029 USA

Raphael Lamed Department of Molecular Microbiology and Biotechnology George S. Wise Faculty of Life Sciences Tel Aviv University Ramat Aviv 69978 Israel

J. Gijs Kuenen Department of Biotechnology Delft University of Technology 2628BC Delft The Netherlands Daisy A. Kuhn Department of Biology California State University Northridge, CA 91330 USA

Giancarlo Lancini Consultant, Vicuron Pharmaceutical 21040 Gerenzano (Varese) Italy Jan W. M. la Rivière Institut für Mikrobiologie Universität Göttingen D-3400 Göttingen Germany

xli

xlii

Contributors

Jared R. Leadbetter Environmental Science and Engineering California Institute of Technology Pasadena, CA 91125-7800 USA Donald J. LeBlanc ID Genomics Pharmacia Corporation Kalamazoo, MI 49001 USA Marc Lecuit Unité des Interactions Bactéries-Cellules INSERM U604 Institut Pasteur 75724 Paris France Adrian Lee School of Microbiology & Immunology University of New South Wales Sydney, New South Wales 2052 Australia Natuschka Lee Lehrstuhl für Mikrobiologie Technische Universität München D-85350 Freising Germany Susan Leschine Department of Microbiology University of Massachusetts Amherst, MA 01003-5720 USA Na (Michael) Li Division of Biostatistics School of Public Health University of Minnesota Minneapolis, MN 55455 USA Mary E. Lidstrom Department of Chemical Engineering University of Washington Seattle, WA 98195 USA

Puspita Lisdiyanti Laboratory of General and Applied Microbiology Department of Applied Biology and Chemistry Faculty of Applied Bioscience Tokyo University of Agriculture Tokyo, Japan Derek Lovley Department of Microbiology University of Massachusetts Amherst, MA 01003 USA Wolfgang Ludwig Lehrstuhl für Mikrobiologie Technische Universität München D-85350 Freising Germany David Lyerly TechLab, Inc. Corporate Research Center Blacksburg VA 24060-6364 USA Janet I. Macinnes University of Guelph Guelph N1G 2W1 Canada Michael T. Madigan Department of Microbiology Mailcode 6508 Southern Illinois University Carbondale, IL 62901-4399 USA Luis Angel Maldonado School of Biology Universidad Nacional Autonoma de Mexico (UNAM) Instituto de Ciencias del Mar y Limnologia Ciudad Universitaria CP 04510 Mexico DF Mexico

Wolfgang Liebl Institut für Mikrobiologie und Genetik Georg-August-Universität D-37077 Göttingen Germany

Jim Manos The University of Maryland Biotechnology Institute Center of Marine Biotechnology Baltimore, MD 21202

Franz Lingens Institut fur Mikrobiologie Universitat Hohenheim D-7000 Stuttgart 70 Germany

Lynn Margulis Department of Botany University of Massachusetts Amherst, MA 01003 USA

Contributors

xliii

Kevin C. Marshall School of Microbiology University of New South Wales Kensington New South Wales 2033 Australia

Ulrich Melcher Department of Biochemistry and Molecular Biology Oklahoma State University Stillwater, OK USA

Esperanza Martinez-Romero Centro de Investigacion sobre Fijacion de Nitrogeno Cuernavaca Mor Mexico

Jianghong Meng Nutrition and Food Science University of Maryland College Park, MD 20742-7521 USA

Vitor A. P. Martins dos Santos Gesellschaft für Biotechnologische Forschung Division of Microbiology Braunschweig D-38124 Germany

Valery V. Mikhailov Pacific Institute of Bioorganic Chemistry Far-Eastern Branch of the Russian Academy of Sciences 690022 Vladivostok Russia

Vega Masignani IRIS, Chiron SpA 53100 Siena Italy Paola Mattarelli Istituto di Microbiologia Agraria 40126 Bologna Italy Carola Matthies Department of Ecological Microbiology BITOEK, University of Bayreuth D-95440 Bayreuth Germany Ann G. Matthysse Department of Biology University of North Carolina Chapel Hill, NC 27599 USA Megan E. McBee Biological Engineering Division Massachusetts Institute of Technology Cambridge, MA USA Bruce A. McClane Department of Molecular Genetics and Biochemistry University of Pittsburgh School of Medicine Pittsburgh, PA 15261 USA Zoe P. McKiness Department of Organic and Evolutionary Biology Harvard University Cambridge, MA 02138 USA

Melissa B. Miller, Ph.D. Department of Pathology and Laboratory Medicine University of North Carolina at Chapel Hill Chapel Hill, NC 27599 USA Michael F. Minnick Division of Biological Sciences University of Montana Missoula, MT 59812-4824 USA Ralph Mitchell Laboratory of Microbial Ecology Division of Engineering and Applied Sciences Harvard University Cambridge, MA 02138 USA Cesare Montecucco Professor of General Pathology Venetian Institute for Molecular Medicine 35129 Padova Italy Edward R. B. Moore The Macaulay Institute Environmental Sciences Group Aberdeen AB158QH UK and Culture Collection University of Göteborg (CCUG) Department of Clinical Bacteriology University of Göteborg Göteborg SE-416 43 Sweden

xliv

Contributors

Nancy A. Moran University of Arizona Department of Ecology and Evolutionary Biology Tucson, AZ 85721 USA

Klaus Neuhaus Department of Pediatrics, Infection, Immunity, and Infectious Diseases Unit Washington University School of Medicine St. Louis, MO 63110 USA

Edgardo Moreno Tropical Disease Research Program (PIET) Veterinary School, Universidad Nacional Costa Rica

Diane G. Newell Veterinary Laboratory Agency (Weybridge) Addlestone New Haw Surrey KT1 53NB UK

Ignacio Moriyón Department of Microbiology University of Navarra 32080 Pamplona Spain Annette Moter Institut für Mikrobiologie und Hygiene Universitaetsklinikum Chariteacute Humboldt-Universität zu Berlin D-10117 Berlin Germany Eppe Gerke Mulder Laboratory of Microbiology Agricultural University 6703 CT Wageningen The Netherlands Yasuyoshi Nakagawa Biological Resource Center (NBRC) Department of Biotechnology National Institute of Technology and Evaluation Chiba 292-0818 Japan Francis Nano Department of Biochemistry & Microbiology University of Victoria Victoria V8W 3PG Canada

Irene L. G. Newton Department of Organismic and Evolutionary Biology Harvard University Cambridge, MA 02138 USA S.A. Nierzwicki-Bauer Department of Biology Rensselaer Polytechnic Institute Troy, NY USA M. Fernanda Nobre Departamento de Zoologia Universidade de Coimbra 3004-517 Coimbra Portugal Philippe Normand Laboratoire d’Ecologie Microbienne UMR CNRS 5557 Université Claude-Bernard Lyon 1 69622 Villeurbanne France Steven J. Norris Department of Pathology and Laboratory Medicine and Microbiology and Molecular Genetics University of Texas Medical Scvhool at Houston Houston, TX 77225 USA

Kenneth H. Nealson Department of Earth Sciences University of Southern California Los Angeles, CA 90033 USA

Howard Ochman Department of Biochemistry and Molecular Biophysics University of Arizona Tucson, AZ 85721 USA

Douglas C. Nelson Department of Microbiology University of California, Davis Davis, CA 95616 USA

Gary E. Oertli Molecular and Cellular Biology Unviersity of Washington Seattle, WA 98195-7275 USA

Contributors

Itzhak Ofek Department of Human Microbiology Tel Aviv University 69978 Ramat Aviv Israel Bernard Ollivier Laboratoire ORSTOM de Microbiologie des Anaérobies Université de Provence CESB-ESIL 13288 Marseille France Scott L. O’Neill Department of Epidemiology and Public Health Yale University School of Medicine New Haven, CT 06520-8034 USA Aharon Oren Division of Microbial and Molecular Ecology The Institute of Life Sciences and Moshe Shilo Minerva Center for Marine Biogeochemistry The Hebrew University of Jerusalem 91904 Jerusalem Israel Jani L. O’Rourke School of Microbiology and Immunology University of New South Wales Sydney, NSW 2052 Australia Jörg Overmann Bereich Mikrobiologie Department Biologie I Ludwig-Maximilians-Universität München D-80638 München Germany Norman R. Pace Department of Molecular, Cellular and Developmental Biology Unversity of Colorado Boulder, CO 80309-0347 USA Norberto J. Palleroni Rutgers University Department of Biochemistry and Microbiology New Brunswick 08901-8525 New Jersey USA

xlv

Bruce Panilaitis Department of Chemcial and Biomedical Engineering Tufts University Medford, MA 02155 USA Nickolas Panopoulos Department of Plant Pathology University of California-Berkeley Berkeley, CA 94720 USA Yong-Ha Park Korean Collection for Type Cultures Korea Research Institute of Bioscience & Biotechnology Taejon 305-600 Korea Matthew R. Parsek University of Iowa Iowa City, IA 52242 USA Bruce J. Paster Department of Molecular Genetics The Forsyth Institute Boston, MA 02115 USA Jerome J. Perry 3125 Eton Road Raleigh, NC 27608-1113 USA Jeannette Pham The CDS Users Group Department of Microbiology South Eastern Area Laboratory Services The Prince of Wales Hospital Campus Randwick NSW 2031 Australia Harvey M. Pickrum Proctor and Gamble Company Miami Valley Laboratories Cincinnatti, OH 45239 USA Dietmar H. Pieper Gesellschaft für Biotechnologische Forschung Division of Microbiology Braunschweig D-38124 Germany Beverly K. Pierson Biology Department University of Puget Sound Tacoma, WA 98416 USA

xlvi

Contributors

Mariagrazia Pizza IRIS, Chiron SpA 53100 Siena Italy Jeanne S. Poindexter Department of Biological Sciences Barnard College/Columbia University New York, NY 10027-6598 USA Andreas Pommerening-Röser Institut für Allgemeine Botanik Abteilung Mikrobiologie Universität Hamburg D-22069 Hamburg Germany Michel R. Popoff Unité des Toxines Microbiennes Institut Pasteur 75724 Paris France Anton F. Post Department of Plant and Environmental Sciences Life Sciences Institute Hebrew University Givat Ram 91906 Jerusalem Israel Bruno Pot Laboratorium voor Microbiologie en Microbiële Genetica Rijksuniversiteit Ghent B-9000 Ghent Belgium David Prangishvili Department of Mikrobiology Universitity of Regensburg D-93053 Regensburg Germany Helmut Prauser DSMZ-German Collection of Microorganisms and Cell Cultures GmbH D-38124 Braunschweig Germany Michael Prentice Bart’s and the London School of Medicine and Dentistry Department of Medical Microbiology St. Bartholomew’s Hospital London EC1A 7BE UK

Ulrike Purkhold Lehrstuhl für Mikrobiologie Technische Universität München D-80290 Munich Germany Wim J. Quax Department of Pharmaceutical Biology University of Groningen Groningen 9713AV The Netherlands Erika Teresa Quintana School of Biology Universtiy of Newcastle Newcastle upon Tyne NE1 7RU UK Ralf Rabus Max-Planck-Institut für Marine Mikrobiologie D-28359 Bremen Germany Reinhard Rachel Lehrstuhl für Mikrobiologie Universität Regensburg D-93053 Regensburg Germany A. N. Rai Biochemistry Department North-Eastern Hill University Shillong 793022 India Frederick A. Rainey Department of Biological Sciences Louisiana State University Baton Rouge, LA 70803 USA Juan-Luis Ramos Estación Experimental del Zaidin Department of Biochemistry and Molecular and Cell Biology of Plants Granada E-18008 Spain Rino Rappuoli IRIS Chiron Biocine Immunobiologie Research Institute Siena 53100 Siena Italy Shmuel Razin Department of Membrane and Ultrastructure Research The Hebrew University-Hadassah Medical School Jerusalem 91120

Contributors

Annette C. Reboli Department of Medicine Hahneman University Hospital Philadelphia, PA 19102 USA David W. Reed Biotechnology Department Idaho National Engineering and Environmental Laboratory (INEEL) Idaho Falls, ID 83415-2203 USA Hans Reichenbach GBF D-3300 Braunschweig Germany Barbara Reinhold-Hurek Laboratory of General Microbiology Universität Bremen Laboratorium für Allgemeine Mikrobiologie D-28334 Bremen Germany Markus Riegler Integrative Biology School University of Queensland Australia Monica Riley Marine Biological Lab Woods Hole, MA 02543 USA Lesley A. Robertson Department of Biotechnology Delft University of Technology 2628 BC Delft The Netherlands Edson R. Rocha Department of Microbiology and Immunology East Carolina University Greenville, NC 27858-4354 USA Palmer Rogers Department of Microbiology University of Minnesota Medical School Minneapolis, MN 55455 USA Lyudmila A. Romanenko Senior Researcher in Biology Laboratory of Microbiology Pacific Institute of Bioorganic Chemistry of the Far-Eastern Branch of the Russian Academy of Sciences Vladivostoku, 159 Russia

xlvii

M. F. Romine Pacific Northwest National Laboratory Richland, WA 99352 USA Eliora Z. Ron Department of Molecular Microbiology and Biotechnology The George S. Wise Faculty of Life Sciences Tel Aviv University Ramat Aviv 69978 Tel Aviv Israel Julian I. Rood Australian Bacterial Pathogenesis Program Department of Microbiology Monash University Victoria 3800 Australia Eugene Rosenberg Department of Molecular Microbiology & Biotechnology Tel Aviv University Ramat Aviv 69978 Tel Aviv Israel Frank Rosenzweig Division of Biological Sciences University of Montana Missoula, MT 59812-4824 USA Ornella Rossetto Centro CNR Biomembrane and Dipartimento di Scienze Biomediche 35100 Padova Italy Michael J. Sadowsky Department of Soil, Water, and Climate University of Minnesota Minneapolis, MN 55455 USA Hermann Sahm Institut Biotechnologie Forschungszentrum Jülich D-52425 Jülich Germany Joseph W. St. Gemer, III Department of Molecular Microbiology Washington University School of Medicine St. Louis, MO 63110 USA

xlviii

Contributors

James E. Samuel Department of Medical Microbiology and Immunology College of Medicine Texas A&M University System Health Science Center College Station, TX, 77843-1114 USA Philippe J. Sansonetti Unité de Pathogénie Microbienne Moléculaire Institut Pasteur 75724 Paris France Scott R. Santos Department of Biochemistry & Molecular Biophysics University of Arizona Tucson, AZ 85721 USA Beatrice Saviola Departments of Molecular Microbiology and Immunology Johns Hopkins School of Hygiene and Public Health Baltimore, MD 21205-2105 USA Klaus P. Schaal Institut für Medizinische Mikrobiologie und Immunologie Universität Bonn D-53105 Bonn Germany David B. Schauer Biological Engineering Division and Division of Comparative Medicine Massachusetts Institute of Technology Cambridge, MA 02139 USA Siegfried Scherer Department für Biowißenschaftliche Grundlagen Wißenschaftszentrum Weihenstephan Technische Universität München D-85354 Freising, Germany Bernhard Schink Fakultät für Biologie der Universität Konstanz D-78434 Konstanz Germany

Hans G. Schlegel Institut für Mikrobiologie der Gessellschaft für Strahlen- und Umweltforschung mbH Göttingen Germany Karl-Heinz Schleifer Lehrstruhl für Mikrobiologie Technische Universität München D-85354 Freising Germany Heinz Schlesner Institut für Allgemeine Mikrobiologie Christian Albrechts Universität D-24118 Kiel Germany Michael Schmid GSF-Forschungszentrum für Umwelt und Gesundheit GmbH Institut für Bodenökologie D-85764 Neuherberg Germany Jean M. Schmidt Department of Botany and Microbiology Arizona State University Tempe, AZ 85287 USA Karin Schmidt Institut für Mikrobiologie Georg-August-Universität D-3400 Göttingen Germany Ruth A. Schmitz University of Göttingen D-3400 Göttingen Germany Hildgund Schrempf FB Biologie/Chemie Universität Osnabrück 49069 Osnabrück Germany Milton N. Schroth Department of Plant Pathology University of California-Berkeley Berkeley, CA 94720 USA Heide N. Schulz Institute for Microbiology University of Hannover D-30167 Hannover Germany

Contributors

Peter Schumann DSMZ-German Collection of Microorganisms and Cell Cultures GmbH D-38124 Braunschweig Germany Arthur Schüßler Institut Botany 64287 Darmstadt Germany Edward Schwartz Institut für Biologie/Mikrobiologie Homboldt-Universität zu Berlin D-10115 Berlin Germany James Scott Geophysical Laboratory Carnegie Institution of Washington Washington, DC 20015 USA Margrethe Haugge Serres Marine Biological Lab Woods Hole, MA 02543 USA James P. Shapleigh Department of Microbiology Cornell University Wing Hall Ithaca, NY 14853-8101 USA Nathan Sharon The Weizmann Institute of Science Department of Biological Chemistry IL-76100 Rehovoth Israel Lawrence J. Shimkets Department of Microbiology The University of Georgia Athens, GA 30602-2605 USA

xlix

Jörg Simon Johann Wolfgang Goethe-Universität Frankfurt Campus Riedberg Institute of Molecular Biosciences Molecular Microbiology and Bioenergetics D-60439 Frankfurt Germany Michel Simonet Départment de Pathogenèse des Maladies Infectieuses et Parasitaires Institut de Biologie de Lille 59021 Lille France Mikael Skurnik Department of Medical Biochemistry University of Turku 20520 Turku Finland James M. Slauch Department of Microbiology College of Medicine University of Illinois and Chemical and Life Sciences Laboratory Urbana, IL 61801 USA Ralph A. Slepecky Department of Biology Syracuse University Syracuse, NY 13244 USA C. Jeffrey Smith Department of Microbiology and Immunology East Carolina University Greenville, NC 27858-4354 USA

Thomas M. Shinnick Center for Infectious Diseases Centers for Disease Control Atlanta, GA 30333 USA

Martin Sobierj Department of Biology Environmental Science and Engineering Oregon Graduate Institute of Science and Technology Portland State University Portland, OR 97291-1000 USA

Yuval Shoham Department of Food Engineering and Biotechnology Technion—Israel Institute of Technology Haifa 32000 Israel

Pamela A. Sokol Department of Microbiology and Infectious Diseases University of Calgary Health Science Center Calgary T2N 4N1 Canada

l

Contributors

Jay V. Solnick Department of Interanl Medicine (Infectious Diseases) and Medical Microbiology and Immunology University of California, Davis School of Medicine Davis, CA 95616 USA Dimitry Yu. Sorokin Department of Biotechnology Delft University of Technology 2628 BC Delft The Netherlands and S.N. Winogradsky Institute of Microbiology 117811 Moscow Russia Georg A. Sprenger Institut Biotechnologie Forschungszentrum Jülich D-52425 Jülich Germany Stefan Spring Deutsche Sammlung von Mikroorganismen und Zellkulturen D-38124 Braunschweig Germany Erko Stackebrandt Deutsche Sammlung von Mikroorganismen und Zellkulturen D-38124 Braunschweig Germany David A. Stahl University of Washington Seattle, WA USA Donald P. Stahly Department of Microbiology University of Iowa Iowa City, IA 52242 USA

Thaddeus B. Stanton PHFSED Research Unit National Animal Disease Center USDA-ARS Ames, IA 50010 USA Daniel C. Stein Department of Cell Biology and Molecular Genetics University of Maryland College Park, MD 20742 USA Reinhard Sterner Universitaet Regensburg Institut fuer Biophysik und Physikalische Biochemie D-93053 Regensburg Germany Karl O. Stetter Lehrstuhl für Mikrobiologie Universität Regensburg D-93053 Regensburg Germany Frank J. Stewart Department of Organic and Evolutionary Biology Harvard University Cambridge, MA 02138 USA Andreas Stolz Institut für Mikrobiologie Universität Stuttgart 70569 Stuttgart Germany Daphne L. Stoner Idaho National Engineering Laboratory EG & G Idaho Idaho Falls, ID 83415 USA

James T. Staley Department of Microbiology University of Washington Seattle, WA 98105 USA

Paul Stoodley Center for Biofilm Engineering Montana State University Bozeman, MT 59717-3980 USA

Alfons J.M. Stams Laboratorium voor Microbiologie Wageningen University NL-6703 CT Wageningen The Netherlands

James R. Swafford Department of Botany and Microbiology Arizona State University Tempe, AZ 85287 USA

Contributors

Jean Swings Laboratorium voor Microbiologie Department of Biochemistry Physiology and Microbiology BCCM/LMG Bacteria Collection Universiteit Gent Gent Belgium Mariko Takeuchi Institute for Fermentation Osaka 532-8686 Japan Ralph Tanner University of Oklahoma Norman, OK, 73019-0390 USA Andreas Teske Department of Marine Sciences University of North Carolina at Chapel Hill Chapel Hill, NC 27599 USA Michael Teuber ETH-Zentrum Lab Food Microbiology CH-8092 Zürich Switzerland Gabriele Timmermann Institut für Allgemeine Botanik Abteilung Mikrobiologie Universität Hamburg D-22069 Hamburg Germany Brian J. Tindall Deutsche Sammlung von Mikroorganismen und Zellkulturen Braunschweig D-38124 Germany Kevin Towner Consultant Clinical Scientist Public Health Laboratory University Hospital Nottingham NG7 2UH UK Hans G. Trüper Institut für Mikrobiologie und Biotechnologie D-53115 Bonn Germany Elaine Tuomanen Department of Infectious Diseases St. Jude Children’s Research Hospital Memphis, TN 38105-2394 USA

Francisco A. Uzal California Animal Health and Food Safety Laboratory University of California, Davis San Bernardino, CA 92408 USA Peter Van damme Laboraroorium voor Microbiologie Faculteit Wetenschappen Universiteit Gent B-9000 Gent Belgium Antonio Ventosa Department of Microbiology and Parasitology Faculty of Pharmacy University of Sevilla 41012 Sevilla Spain Gernot Vobis Centro Regional Universitario Bariloche Universidad Nacional de Comahue Barioloche 8400, Rio Negro Argentina Alexander von Graevenitz Department of Medical Microbiology University of Zürich GH-8028 Zürich Switzerland Günther Wächtershäuser 80331 Munich Germany Lawrence P. Wackett Department of Biochemistry, Molecular Biology and Biophysics and Biological Process Technology Institute University of Minnesota St. Paul, MN, 55108-1030 USA William G. Wade Department of Microbiology Guy’s Campus London, SE1 9RT UK Michael Wagner Lehrstuhl für Mikrobielle Ökologie Institut für Ökologie und Naturschutz Universität Wien A-1090 Vienna Austria

li

lii

Contributors

David H. Walker Department of Pathology University of Texas Medical Branch Galveston, TX 77555-0609 USA

Jürgen Wiegel University of Georgia Department of Microbiology Athens, GA 30602 USA

Naomi Ward The Institute for Genomic Research Rockville, MD 20850 USA

Robert A. Whiley Queen Mary, University of London London E1 4NS UK

Trudy M. Wassenaar Molecular Microbiology and Genomics Consultants 55576 Zotzenheim Germany

Tracy Whilkins TechLab, Inc. Corporate Research Center Blacksburg VA 24060-6364 USA

John B. Waterbury Woods Hole Oceanographic Institution Woods Hole, MA 02543 USA

Anne Willems Laboratorium voor Mikrobiologie Universiteit Gent B-9000 Gent Belgium

Georges Wauters Université Catholique de Louvain Faculté de Médecine Unité de Microbiologie B-1200 Bruxelles Belgium

Carl R. Woese Department of Microbiology University of Illinois Urbana, IL 61801 USA

Astri Wayadande Department of Entomology and Plant Pathology Oklahoma State University Stillwater, OK USA Alison Weiss Molecular Genetics, Biology and Microbiology University of Cincinnati Cincinnati, OH 45267 USA Rodney A. Welch Medical Microbiology and Immunology University of Wisconsin Madison, WI 53706-1532 USA

Ralph S. Wolfe Department of Microbiology University of Illinois Urbana, IL 61801 Ann P. Wood Division of Life Sciences King’s College London London WC2R 2LS UK Donald E. Woods Department of Microbiology and Infectious Diseases University of Calgary Health Science Center Calgary T2N 4N1 Canada

William B. Whitman Department of Microbiology University of Georgia Athens, GA 30605-2605 USA

B. W. Wren Department of Infectious and Tropical Diseases London School of Hygiene and Tropical Medicine London WC1E 7HT UK

Friedrich Widdel Max-Planck-Institut für Marine Mikrobiologie D-28359 Bremen Germany

Timothy L. Yahr University of Iowa Iowa City, IA 52242 USA

Contributors

Atteyet F. Yassin Institut für Medizinische Mikrobiologie und Immunologie Universität Bonn D-53105 Bonn Germany Jung-Hoon Yoon Korean Collection for Type Cultures Korea Research Institute of Bioscience and Biotechnology Yuson, Taejon 305-600 Korea Allan A. Yousten Biology Department Virginia Polytechnic Institute and State University Blacksburg, VA 24061 USA Xue-Jie Yu University of Texas Medical Branch Galveston, TX USA

Vladimir V. Yurkov Department of Microbiology University of Manitoba Winnipeg R3T 2N2 Canada Georgi A. Zavarzin Institute of Microbiology Academy of Sciences of the USSR 117312 Moscow Russia Mary Jo Zidwick Cargill Biotechnology Development Center Freshwater Building Minneapolis, MN 55440 USA Stephen H. Zinder Department of Microbiology Cornell University 272 Wing Hall Ithaca, NY 14853 USA

liii

Prokaryotes (2006) 5:3–37 DOI: 10.1007/0-387-30745-1_1

no i t cudor tnI

ot

eht

a i re t caboe torP

Introduction to the Proteobacteria KAREL KERSTERS, PAUL DE VOS, MONIQUE GILLIS, JEAN SWINGS, PETER VANDAMME AND ERKO STACKEBRANDT

Introduction Within the domain Bacteria, the phylum Proteobacteria constitutes at present the largest and phenotypically most diverse phylogenetic lineage. In 1988, Stackebrandt et al. named the Proteobacteria after the Greek god Proteus, who could assume many different shapes, to reflect the enormous diversity of morphologies and physiologies observed within this bacterial phylum. In 2002, the Proteobacteria consist of more than 460 genera and more than 1600 species, scattered over 5 major phylogenetic lines of descent known as the classes “Alphaproteobacteria,” “Betaproteobacteria,” “Gammaproteobacteria,” “Deltaproteobacteria” and “Epsilonproteobacteria.” The Proteobacteria account for more than 40% of all validly published prokaryotic genera and encompass a major proportion of the traditional Gramnegative bacteria, show extreme metabolic diversity, and are of great biological importance, as they include the majority of the known Gramnegatives of medical, veterinary, industrial and agricultural interest. Moreover, not only can the origin of mitochondria be traced back to the “Alphaproteobacteria,” but several representatives are ecologically important because they play key roles in the carbon, sulfur and nitrogen cycles on our planet. In this context, the purple nonsulfur bacteria and the rhizobia (both members of the “Alphaproteobacteria”) are two of the most apparent and well-known examples, because the former are purple-colored photosynthetic prokaryotes using light as energy source, and the latter are able to reduce atmospheric nitrogen gas when living in symbiosis with leguminous plants. Agriculture and life on earth would be very different in the absence of these nitrogen-fixing rhizobia! The Proteobacteria, formerly known as “purple bacteria and relatives,” are characterized by a bewildering diversity of morphological and physiological types: besides rods and cocci, curved, spiral, ring-shaped, appendaged, filamentous and sheathed bacteria occur among this phylum. Most Proteobacteria are meso-

philic, but some thermophilic (e.g., Thiomonas thermosulfata and Tepidomonas) and psychrophilic (e.g., Polaromonas) representatives have been described. A great number of Proteobacteria are motile by means of polar or peritrichous flagella, whereas the myxobacteria (belonging to the “Deltaproteobacteria”) display a gliding type of motility and show highly complex developmental lifestyles, whereby often remarkable multicellular and macroscopic structures (so-called “fruiting bodies”) are formed. Most of the known Proteobacteria are free-living; some (such as the rhizobia) enter in symbiotic associations with specific leguminous plants, where they fix nitrogen in root or stem nodules. Others live as intracellular endosymbionts of protozoa and invertebrates (mussels, insects and nematodes), whereas the rickettsiae are obligate intracellular parasites of humans or mammals. The extreme diversity of energygenerating mechanisms is a unique biochemical characteristic of the Proteobacteria: some are chemoorganotrophs (e.g., Escherichia coli), others are chemolithotrophs (e.g., the sulfuroxidizing bacteria such as the thiobacilli and the ammonia-oxidizing bacteria such as Nitrosomonas) or phototrophs (e.g., the purple colored Chromatium, Rhodospirillum and many others). Concerning their relationship towards oxygen, the Proteobacteria include strictly aerobic and anaerobic species as well as facultative aerobes and microaerophiles. Denitrifiers are reported among the “Alphaproteobacteria,” “Betaproteobacteria,” “Gammaproteobacteria,” and the “Epsilonproteobacteria.” To reveal the extreme phenotypic biodiversity among the Proteobacteria, Table 1 gives an overview of the distribution of the five phylogenetic proteobacterial groups among 13 major phenotypic groups as defined in the 9th edition of Bergey’s Manual of Determinative Bacteriology (Holt et al., 1994). Table 1 clearly shows that only three of these 13 phenotypic groups are unique to any one of the current proteobacterial classes. Representatives of some of these 13 phenotypic groups occur also among several other prokaryotic phyla.

4

K. Kersters et al.

Table 1. The occurrence of major phenotypic groups among the five classes of the Proteobacteria. Phenotypic groupa Aerobic/microaerophilic, motile, helical/vibrioid Gram-negative bacteria Gram-negative aerobic/microaerophilic rods and cocci Facultatively anaerobic Gram-negative rods Anaerobic straight, curved, and helical Gram-negative rods Symbiotic and parasitic bacteria of vertebrate and invertebrate species Anoxygenic phototrophic bacteria Aerobic chemolithotrophic bacteria and associated genera Budding and/or appendaged nonphototrophic bacteria Nonphotosynthetic, nonfruiting gliding bacteria Nonmotile or rarely motile, curved Gram-negative bacteria Dissimilatory sulfate- or sulfur-reducing bacteria Sheathed bacteria Fruiting gliding bacteria

Group nr.a

Occurring in proteobacterial class:

2 4 5 6 9 10 12 13 15 3 7 14 16

α, β, γ, δ and ε α, β, γ, δ and ε α, β, γ, δ and ε β, γ and δ α and γ α, β and γ β, γ and ε α and γ β and γ α and ε δ β δ

a

Phenotypic groups and group numbers are according to Bergey’s Manual of Determinative Bacteriology (Holt et al., 1994). See also Garrity and Holt (2001).

This chapter sketches how and why the Proteobacteria were recognized as a coherent but diverse phylogenetic group and summarizes its phylogenetic structure as well as its phenotypic and ecological diversity.

Towards a Phylogenetic Definition of the “Purple Bacteria and Their Nonphototrophic Relatives” In the context of the above mentioned extreme biodiversity of the Proteobacteria, it is not surprising that some 50 years ago—when only phenotypic techniques were available for characterization and comparison of prokaryotes—it was impossible to foresee that morphologically and physiologically highly different bacterial groups such as the Enterobacteriaceae and the photosynthetic purple sulfur bacteria (e.g., Chromatium) would be phylogenetically more closely related to each other, whereas morphologically similar bacteria such as the Enterobacteriaceae and the flavobacteria would harbor widely different genomes. Microbiologists possessed in those years only tools to look at the tips of the evolutionary branches of the tree of life. About 15 years prior to the introduction of ribosomal RNA analysis as a suitable method for determining prokaryote evolution, two different approaches were followed aiming at the same goal. The physiologists used the deductive strategy, trying to establish the phylogeny of prokaryotes from selected metabolic properties (Broda, 1970; Broda, 1975), while geneticists and biochemists followed the inductive strategy, originating from the pioneering publication by Zuckerkandl and Pauling (1965). The question about the origin of bacterial respiration placed the photosynthetic bacteria, especially the anoxygenic “purple bacteria,” in

the center of discussion. To be more precise, the problem was how respiration could evolve in parallel in so many groups of organisms and independently from that evolved in cyanobacteria. The similarity of the electron-flow chains in photosynthesis and respiration as an ordered assembly of flavoproteins, cytochromes, quinones and non-heme Fe-S proteins in connection with membranes has been noted (Olson, 1970), and it was proposed that the photosynthetic chain could have been modified and adapted to respiration. It was Broda (1970) who concluded that respiration evolved independently from different kinds of photosynthetic apparatus, and if photosynthetic bacteria themselves evolved in different lines of descent, so would the aerobic respiring bacteria. This “conversion” hypothesis placed significant emphasis on the presence of phototrophic bacteria in various lines of descent and focused on the evolution of electron chains from photosynthetic anaerobes, anaerobic respirers, and aerobes ranging from phototrophic aerobes to strictly aerobic respirers. Broda’s hypothesis, later supported (though refined) by the general phylogenetic frame of ribosomal RNA/DNA sequences, is illustrated by two examples. Firstly, he postulated the existence of a Gram-positive phototroph, later proven by the isolation of the phylogenetically Gram-positive Heliobacterium chlorum (Gest and Favinger, 1983; Woese et al., 1985a), branching deeply in the Clostridium lineage. Secondly, early 16S rRNA cataloguing data indicated the high phylogenetic similarity between the nonsulfur purple bacterium Rhodopseudomonas palustris and the aerobic Nitrobacter winogradskyi (Seewaldt et al., 1982). However, Broda (1975) was not in a position to predict the specific phylogenetic clustering of the purple sulfur and the purple nonsulfur bacteria as opposed to the lin-

Introduction to the Proteobacteria

eages of green sulfur bacteria, the green nonsulfur bacteria, and the cyanobacteria. Also, as they had not been described in the 1970s, the aerobic bacteriochlorophyll a-containing bacteria (α-3 and α-4 groups and a few in the “Betaproteobacteria”) are not mentioned in the conversion hypothesis. Modifications to the outline of biochemical pathways given by Broda have been made by Schwemmler (1989), combining rRNAbased phylogenetic relatedness between species and the biochemistry of components of electron chains involved in aerobic respiration and anaerobic and aerobic photosynthesis. Zuckerkandl and Pauling (1965) highlighted the importance of using semantides, such as genomic DNA, its primary transcript RNA, and the translation product, the proteins, as evolutionary markers. They postulated that the path of evolution is laid down in the primary structure of nucleic acids and determination of the blueprint of evolutionary conservative genes would necessarily unravel the evolutionary history of organisms. Owing to methodological restrictions, proteins rather than DNA or RNA were accessible to sequence analyses, restricting early molecular studies to those proteins easily isolated and sequenced. Besides histones and fibrinopeptides, used in the determination of the rate of gene evolution in eukaryotes, primarily proteins such as ferredoxin and cytochrome c allowed the first glimpse of the evolution of prokaryotic genes. While the ferredoxin data were obtained for mainly Gram-positive anaerobic bacteria and cyanobacteria, cytochrome c sequences were generated for respiring organisms as well as for anaerobic and aerobic photosynthetic bacteria (Schwartz and Dayhoff, 1978; Ambler et al., 1979) as well as for mitochondria, the origin of which could be traced to members of the “Alphaproteobacteria.” Surprising to taxonomists, the clustering of species according to the primary and tertiary structure of proteins (Dickerson, 1980) did not correlate with the traditional systematic groupings such as those laid down in Bergey’s Manual of Determinative Bacteriology (Buchanan and Gibbons, 1974). It was especially the phylogenetic closeness between Gram-negative, nonphototrophic organisms and phototrophs that shed doubts on the correctness of the sequence-based findings. However, shortly afterwards, results from the 16S rRNA cataloguing approach, fully supporting the protein-based studies, demonstrated the obvious failure of the phenotype to mirror phylogenetic relatedness. Some 30 years ago, the pioneering studies on 16S rRNA oligonucleotide cataloguing by Woese and coworkers (Fox et al., 1977; Woese and Fox, 1977) revolutionized the insights in microbial evolution, in particular, and evolution of life on earth, in general. The purple

5

bacteria (nowadays named Proteobacteria) were first recognized and circumscribed as a distinct division (phylum) of the eubacteria by Woese and coworkers on the basis of signature and sequence analyses of the 16S rRNA/rDNA (Gibson et al., 1979; Fox et al., 1980; Woese et al., 1985b; Woese, 1987). Because the purple photosynthetic phenotype was distributed throughout the group the trivial name “purple bacteria” seemed to be justified for this phylogenetic group of Gram-negatives (Woese, 1987), although many nonphotosynthetic species (such as the enterics, the pseudomonads, rhizobia, rickettsiae, etc.) grouped among the photosynthetic representatives. The oligonucleotide signature analysis of 16S rRNAs revealed already in the 1980s that the purple bacteria comprised at least three major subdivisions, arbitrarily designated as α, β and γ. The purple nonsulfur (PNS) bacteria (e.g., Rhodospirillum, Rhodobacter and Rhodocyclus) were found in two subdivisions (α and β; Woese et al., 1984a; Woese et al., 1984b), whereas the purple sulfur (PS) bacteria (e.g., Chromatium and Ectothiorhodospira) belonged to the γ-subdivision (Woese et al., 1985c). When more Gram-negatives were investigated by rRNA-cataloguing, rDNAsequencing and also by the DNA/rRNA hybridization technique, it became clear that the majority of the so-called purple bacteria were in fact nonphotosynthetic prokaryotes. Without being able to trace the deeper phylogenetic relationships, De Ley and coworkers (reviewed by De Ley, 1992) studied hundreds of Gram-negatives by the DNA/rRNA hybridization technique, which allowed their relative phylogenetic position within the various rRNA-branches to be determined. The α, β and γ groups corresponded to the rRNA superfamilies IV, III and I+II, respectively, as defined by De Ley and coworkers (De Vos and De Ley, 1983; De Vos et al., 1985; De Vos et al., 1989; De Ley et al., 1986; Jarvis et al., 1986; Rossau et al., 1986; Willems et al., 1991a; De Ley, 1992). The DNA/rRNA hybridization approach together with the application of various genomic and phenotypic techniques (i.e., the polyphasic approach; Colwell, 1970; Vandamme et al., 1996) allowed improvement of the classification of various Gram-negative taxa, such as the pseudomonads (De Vos and De Ley, 1983; De Vos et al., 1985; De Vos et al., 1989; Willems et al., 1991a; De Ley, 1992; Kersters et al., 1996), Alcaligenes and Bordetella (Kersters and De Ley, 1984; De Ley et al., 1986), the Pasteurellaceae (De Ley et al., 1990), the Neisseriaceae and Moraxellaceae (Rossau et al., 1986; Rossau et al., 1991), the Acetobacteraceae (Gillis and De Ley, 1980; De Ley and Gillis, 1984; Swings, 1992), the Campylobacteraceae (Vandamme and De Ley, 1991) and many others.

6

K. Kersters et al.

During the last 15 years, the knowledge concerning the phylogeny of the Gram-negatives gradually became more comprehensive by the availability of an increasing number of nearly complete 16S rRNA sequences, indicating that the Gram-negative sulfur- and sulfate-reducing bacteria form a fourth group or δ-subdivision among the purple bacteria together with the myxobacteria and the bdellovibrios (Oyaizu and Woese, 1985; Woese, 1987). Because the majority of the genera belonging to the purple bacteria are not purple and not photosynthetic, Stackebrandt et al. proposed in 1988 the name “Proteobacteria” for a new higher taxon (at the level of class), including these purple bacteria and their relatives; the α to δ groups were temporarily considered as subclasses, pending further studies and nomenclatural proposals. Later some microaerophilic and helical-shaped bacteria such as the campylobacters were placed in the fifth or ε-subclass, corresponding to rRNAsuperfamily VI of De Ley’s research group (Vandamme and De Ley, 1991; Stackebrandt, 1992). Each new issue of the International Journal of Systematic and Evolutionary Microbiology adds new genera and species to the Proteobacteria, entailing a more bush-like rRNA/rDNA-tree topology. Yet, the distinctness of the subclasses seems to be maintained, although the differentiation between the β- and the γ-group is becoming less clear, and some rDNA-based trees indicate that Desulfurella and allied bacteria of the δgroup may constitute a sixth subdivision (Rainey et al., 1993). In the second edition of Bergey’s Manual of Systematic Bacteriology (Garrity, 2001a), the Proteobacteria have been elevated to the rank of phylum and the subclasses α to ε have been elevated to the rank of classes, corresponding to the names “Alphaproteobacteria,” “Betaproteobacteria,” “Gammaproteobacteria,” “Deltaproteobacteria,” and “Epsilonproteobacteria,” respectively (see Table 2). In a recently

revised megaclassification of the prokaryotes, Cavalier-Smith (2002) proposes a new classification and nomenclature for the five major subgroups of the Proteobacteria (see Table 2). In the present chapter, we follow the classification of the major reference work, Bergey’s Manual of Systematic Bacteriology (Garrity, 2001), and will use ranks and names as listed in Table 2 (2nd column). Is the 16S-rRNA-based Phylogeny of the Proteobacteria Confirmed by Other Experimental Approaches? The first indications that the traditional groupings of the Gram-negative bacteria according to the previous editions of Bergey’s Manual of Systematic Bacteriology (Buchanan and Gibbons, 1974; Krieg and Holt, 1984) and Bergey’s Manual of Determinative Bacteriology (Holt et al., 1994) were not phylogenetic came from comparative sequence analysis of some redox proteins, such as cytochrome c [Introduction]. Also studies by Byng et al. (1983) on the regulation of multibranched pathways for the biosynthesis of aromatic amino acids indicated that some members of the genus Pseudomonas, such as P. testosteroni and P. solanacearum, were more similar to other genera than to the group of species around the type species Pseudomonas aeruginosa (belonging to the “Gammaproteobacteria”), which is in perfect agreement with rRNA/ rDNA data. Later, Pseudomonas testosteroni and P. solanacearum (both belonging to the “Betaproteobacteria”) were indeed transferred to the genera Comamonas and Ralstonia, respectively (Tamaoka et al., 1987; Yabuuchi et al., 1995). The definition of the Proteobacteria as a separate evolutionary lineage and the overall picture of the grouping in five classes are confirmed by comparative analysis of 23S rRNA-genes and alternative phylogenetic markers such as the genes coding for elongation factor Tu and

Table 2. Three systems of Proteobacteria classification: correspondence between the names of taxa above the rank of order. Classification 1a

2b

Class Proteobacteria Subclass alpha Subclass beta Subclass gamma Subclass delta Subclass epsilon a

Phylum Proteobacteria Class “Alphaproteobacteria”d Class “Betaproteobacteria” Class “Gammaproteobacteria” Class “Deltaproteobacteria” Class “Epsilonproteobacteria”

From Stackebrandt et al. (1988b). From Bergey’s Manual of Systematic Bacteriology (Garrity, 2001). c From Cavalier-Smith (2002). d Quotation marks are used for names that have not yet been validated. b

3c Division Proteobacteria Subdivision Rhodobacteria Class Alphabacteria

}

Class Chromatibacteria Subdivision Thiobacteria Class Deltabacteria Class Epsilobacteria

Introduction to the Proteobacteria

ATPase (Ludwig et al., 1995; Ludwig and Schleifer, 1999; Ludwig and Klenk, 2001). Moreover, the comparative analysis of conserved insertions and deletions (so-called “signature sequences”) found in different bacterial proteins yielded molecular means to define the phylum Proteobacteria and its various classes and improved the understanding of their evolutionary relationships to other groups of Bacteria and Eukarya (Gupta, 2000; Gupta, 2002). Proteins studied were, e.g., alanyl-tRNA synthetase, succinyl-CoA synthetase, Hsp 60 (GroEL), Hsp70 heat shock protein, and recA protein. The listing of extensive literature on this subject would be unjustifiable in the context of this chapter (see Gupta, 2000). The proteobacterial classes emerged in the following order during evolution (Gupta, 2000): epsilon and delta → alpha → beta → gamma, which is similar to the order proposed on the basis of RNA polymerase β and β′ subunits (Klenk et al., 1999). Indeed, the ribosomal DNA tree does not clearly resolve the order in which the majority of main lineages emerged during evolution. Proteobacteria, Gram-positive bacteria, Cyanobacteria, Cytophaga/Flavobacteria and some other minor lineages appear to evolve at the same node, giving the rDNA-tree a fork-like appearance. The branching order of the various main lines of descent as depicted by Gupta (2000) is different. Gupta places the low G+C Gram-positive bacteria (clostridia) at the base of bacterial evolution, while those organisms constituting the base of the bacterial 16S rDNA tree (Aquifex, Deinococcus-Thermus, and green sulfur bacteria) are described as having evolved later. Proteobacteria appear as the last evolutionary group. The importance of the proteobacterial cell is highlighted not only as the donor of the mitochondria (Margulis, 1993; Falah and Gupta, 1994), but also as one of the fusion partners (the other being a archaeal cell) giving rise to the ancestral eukaryotic cell (see Gupta [2000] for an extensive list of references). More genes and complete genomes will have to be sequenced to determine the extent and relative time of lateral gene transfer and to highlight those genes that represent the core of genes transmitted vertically, hence representing the evolutionarily stable component of the cell. So far, no reliable phenotypic features were found that would be characteristic for the whole phylum of the Proteobacteria and differentiate it from other phyla of the domain Bacteria. It seems also very difficult to find stable phenotypic features for the differentiation of the five classes, which is not surprising in view of the enormous variety of morphological and physiological types existing in each of these five lineages. Phototrophy cannot be used as phylogenetic marker, except that PNS- and PS-bacteria are only

7

reported so far among the Proteobacteria, which could indicate that this type of photosynthesis was common in the proteobacterial ancestors and was lost during evolution in a great number of sublineages. Support for the validity of some of the results of nucleic acid and protein sequencing came from chemotaxonomy. Within the “Alphaproteobacteria,” Rhodopseudomonas palustris, Nitrobacter winogradskyi, Blastochloris (Rhodopseudomonas) viridis, Phenylobacterium immobile, certain thiobacilli, Brucella melitensis and Brevundimonas (Pseudomonas) diminuta exhibited an unusual lipid A, which lacked glucosamine but contained instead a 2,3-diamino2,3-dideoxy-D-glucose (Stackebrandt et al., 1988a). This structure was not uniformly distributed among all members of the “Alphaproteobacteria” but was shared among closely related species. Lipid A composition also confirmed the phylogenetic distinctness of species hitherto affiliated to the same genus on the basis of their phenotype. For example, Rhodocyclus tenue and Rhodocyclus gelatinosa (members of the “Betaproteobacteria”; Imhoff et al., 1984) differed significantly in the chemistry of their lipid A, a finding that supported the reclassification of the latter species as Rubrivivax gelatinosus (Willems et al., 1991b). As far as we could determine from the literature, quinones and polyamines seem to be good marker molecules to differentiate the major proteobacterial lineages (Collins and Jones, 1981; Collins and Widdel, 1986; Busse and Auling, 1988; Auling, 1992; Hamana and Matsuzaki, 1993; Busse et al., 1996; Table 3; Directory, Databases and Dictionaries Compiled by WDCM website (http://www.wdcm. nig.ac.jp/cgi-bin/search.cgi)). Ubiquinones are typical for the α, β and γ lineages, whereas menaquinones are characteristic for the “Deltaproteobacteria” and “Epsilonproteobacteria,” although some photosynthetic “Alphaproteobacteria” and “Betaproteobacteria” contain also MK-8, MK-9 or MK-10 (Hiraishi et al., 1984). Some members of the Pasteurellaceae (“Gammaproteobacteria”) are characterized by demethylmenaquinones. Gas chromatographic analysis of the methylesters of cellular fatty acids (FAME) does not allow differentiation of the five major lineages. Of course, phenotypic and chemotaxonomic features (e.g., FAME) are very useful for the description and differentiation of species, genera and families, and the polyphasic taxonomic approach integrating all available data (Colwell, 1970; Vandamme et al., 1996) has been applied during the last decade to improve the classification of previously polyphyletic proteobacterial genera, such as Erwinia (Hauben et al., 1998), Pseudomonas (Kersters et al., 1996; Anzai et al., 2000), Rhizobium and allies (de Lajudie et al., 1998), Thiobacillus (Kelly and

8

K. Kersters et al.

Table 3. Some selected key genera, general characteristics, and differentiating features of the five classes of the Proteobacteria. Proteobacterial class Alpha

Beta

Gamma

Delta

Epsilon

Important genera

Acetobacter Agrobacteriuma Bartonellaa Bradyrhizobium Brucellaa Caulobactera Ehrlichia Gluconobacter Hyphomicrobium Mesorhizobiuma Methylobacteriumb Nitrobacter Rhizobium Rhodobacterb Rhodospirillum Sinorhizobiuma Sphingomonasb Rickettsiaa,b Wolbachiab

Alcaligenes Bordetellaa,b Burkholderiab Comamonas Neisseriaa,b Nitrosomonasb Ralstoniab Rhodocyclus Sphaerotilus Spirillum Thiobacillus

Actinobacillusb Azotobacter Buchneraa Chromatium Coxiellab Erwiniab Escherichiaa,b Francisellab Haemophilusa,b Legionellab Methylococcusb Pasteurellaa Pectobacterium Pseudomonasa,b Salmonellaa,b Shewanellab Shigellaa,b Stenotrophomonas Vibrioa,b Xanthomonasa,b Xylellaa,b Yersiniaa,b

Bdellovibrio Chondromyces Desulfobacter Desulfovibriob Geobacterb Myxococcusb Polyangium Syntrophus

Campylobactera Helicobactera Sulfurospirillum Wolinella

Number of genera/number of speciesc

140/425

76/225

181/755

57/165

6/49

Major ubiquinone typed

Q-10

Q-8

Q-8, Q-9, or Q-10 to Q-14

—

—

Major menaquinone typed

Some contain also MK-9 or MK-10

Some contain also MK-8

Some contain also MK-8 or MK-7

MK-6, MK6(H2), MK7, MK-7(H2) or MK-8e

MK-6, methylsubstituted MK-6f

Characteristic polyaminesg

Most contain a triamine (symhomospermidine or spermidine)

2-Hydroxyputrescine

Spermidine and/or putrescine or cadaverine; or 1,3-diaminopropane

Most contain a triamine (symhomospermidine or spermidine)

Spermidine

Symbols and abbreviations: —, absent; DMK, demethylmenaquinone; and MK-6(H 2), hydrogenated menaquinone-6. a The genome of at least one representative strain has been sequenced (as of mid 2002). b Sequencing of the genome of at least one representative strain is in progress (as of mid 2002). c Only validly published names (situation as of mid 2002). d Collins and Jones (1981), Hiraishi et al. (1984), (http://www.wdcm.nig.ac.jp/cgi-bin/search.cgi), and H.J. Busse, personal communication. e Collins and Widdel (1986). f Moss et al. (1990). g Auling (1992), Busse and Auling (1988), and Hamana and Matsuzaki (1993).

Wood, 2000), Leptothrix (Spring et al., 1996) and Oceanospirillum (Satomi et al., 2002). No single nucleotide signature is found in the 16S rRNA that could serve unambiguously to define the proteobacterial phylum (sensu Bergey’s Manual). However, specific 16S rRNA sequence signatures for the various classes of the

Proteobacteria have been described and used for the construction of DNA probes (Woese, 1987; Stackebrandt et al., 1988b; Manz et al., 1992; Ludwig et al., 1998). Such probes were extensively applied for the detection and visualization of Proteobacteria and other prokaryotes in activated sludge (Wagner et al., 1993; Snaidr et al.,

Introduction to the Proteobacteria

9

Table 4. Some rRNA-targeted oligonucleotide probes for fluorescent in-situ hybridization. Probe

Position

ALF1b

16S rRNA 19–35 23S rRNA 1027–1043 23S rRNA 1027–1043 16S rDNA 385–402

BET42a GAM42a Delta 385

Probe sequence (5′ → 3′) CGTTCG(C/T)TCTGAGCCAG GCCTTCCCACTTCGTTT GCCTTCCCACATCGTTT CGGCGT(C/T)GCTGCGTCAGG

1997; Juretschko et al., 2002). Table 4 lists the sequence of some representative rRNA-targeted oligonucleotide probes.

The Proteobacterial Classes: Morphological, Physiological, Ecological and Phylogenetic Diversity Figure 1 is a simplified phylogenetic tree of the Proteobacteria based on the nearly complete 16S rDNA sequences of the type strains of the type species of the majority of proteobacterial genera. Only the names of the families and major groups are indicated. The “Deltaproteobacteria” and “Epsilonproteobacteria” form the deeper branches of the phylum; the “Alphaproteobacteria” are also clearly separated, whereas the closer relationship between the β and γ lineages may indicate the common origin of the latter groups (Ludwig and Klenk, 2001). Phototrophic bacteria occur only in the “Alphaproteobacteria,” “Betaproteobacteria” and “Gammaproteobacteria” (purple colored triangles in Fig. 1); these anoxygenic phototrophic purple bacteria can be subdivided into the purple sulfur (PS; e.g., Chromatium and Ectothiorhodospira) and purple nonsulfur (PNS) bacteria (e.g., Rhodospirillum). All known PS bacteria belong to the “Gammaproteobacteria” and use H2S or S° as sole electron donor. The PNS bacteria occur among the “Alphaproteobacteria” and “Betaproteobacteria.” A great number of phototrophic Proteobacteria are versatile and can easily switch from a phototrophic to a heterotrophic lifestyle in the absence of light. The results of comparative 16S rDNA sequence analysis have led to extensive taxonomic rearrangements within previously defined taxa of proteobacterial phototrophs (Imhoff, 2001a). In this chapter, we briefly touch on the morphological, physiological, ecological and phylogenetic diversity of the major proteobacterial groups. The reader will find more detailed infor-

Specificity

Reference

“Alphaproteobacteria,” but not exclusive “Betaproteobacteria”

Manz et al., 1992

“Gammaproteobacteria,” but not the deeply branching taxa “Deltaproteobacteria” sulfatereducers, but not exclusive

Manz et al., 1992 Manz et al., 1992 Rabus et al., 1996

mation in the chapters dealing with the individual families and genera of the Proteobacteria. An incomplete survey of disease-causing Proteobacteria is given in Table 5 for the bacteria of clinical and veterinary interest and in Table 6 for the phytopathogens. Some examples of industrial and biotechnological applications are listed in Table 7.

The “Alphaproteobacteria” The 16S rDNA-tree separates the α-class clearly from the other proteobacterial classes (Fig. 1). The bacterial taxa belonging to the “Alphaproteobacteria” (some140 genera and 425 species at present) are morphologically and metabolically extremely diverse. More detailed information can be found in the chapters dealing with the individual families and genera of this class. Table 8 provides an overview of the major phylogenetic groups (see also Fig. 1) and the names of the orders and families of the “Alphaproteobacteria” as they are listed in the 2nd edition of Bergey’s Manual of Systematic Bacteriology (Garrity, 2001a). The majority of the “Alphaproteobacteria” are rod-shaped, but cocci and curved, spiral, stalked, budding and prosthecate forms do also occur. Some are phototrophic purple nonsulfur (PNS) bacteria (such as Rhodospirillum and Rhodobacter), whereas others are chemolithotrophs (e.g., the nitrite-oxidizing Nitrobacter) or chemoorganotrophs (e.g., Sphingomonas and Brucella). Marine and halophilic phototrophic PNS bacteria seem to be restricted to the “Alphaproteobacteria” (Imhoff, 2001b), whereas a physiologically remarkable group of bacteria, containing bacteriochlorophyll (BChl) but unable to grow phototrophically under anaerobic conditions, belongs to various lineages of the “Alphaproteobacteria” (Yurkov and Beatty, 1998; Yurkov, 2001). Except for Roseateles, all aerobic BChl-containing bacteria investigated so far are exclusively found within the “Alphaproteobacteria” and appear to be related phylogenetically to aerobic purely chemoorganotrophic bacteria. This could indicate that the presence of

10% Acetobacteraceae Ehnliciaceae, Rickettsiaceae Holospora group Rhodospirillaceae Sphingomonadaceae

Rhodobacter group

“ALPHAPROTEOBACTERIA”

Caulobacteraceae Rhizobiaceae, Bartonellaceae, Brucellaceae Phyllobacterium group Hyphomicrobiaceae Bradyrhizobium group Methylobacterium, Methylocystis, Rhodobium groups Neisseriaceae Rhodocyclus group Hydrogenophilus group Methylophilus, Nitrosomonas groups Burkholderia, Oxalobacter groups Ralstoria group

“BETAPROTEOBACTERIA”

Alcaligenaceae Comamonadaceae

Xanthomonas group

Enterobacteriaceae, Aeromonadaceae, Alteromonadaceae, Pasteurellaceae, Succinivibrionaceae, Vibrionaceae

Francisella, Piscirickettsia groups Cardiobacteriaceae Moraxellaceae Methylococcaceae Thiothrix group Halomonadaceae Pseudomonadaceae Oceanospirillum group

“GAMMAPROTEOBACTERIA”

Chromatiaceae Legionellaceae, Coxiella group Ectothiorhodospiraceae Myxococcaceae, Archangiaceae, Cystobacteraceae Polyangiaceae Bdellovibrio group Desulfovibrio, Desulfomicrobium groups Desulfohalobium group Desulfuromonas, Geobacter, Pelobacter groups Desulfobacter group Desulfobulbus group Synthrophus group Desulfomonile group Synthrophobacter group Desulfurella group Campylobacteraceae Helicobacter group

“DELTAPROTEOBACTERIA”

“EPSILONPROTEOBACTERIA”

Fig. 1. Simplified neigbor-joining phylogenetic tree of the Proteobacteria based on the 16S rDNA sequences of the type strains of the proteobacterial genera. Distances were calculated using the substitution rate calibration method in TREECON 3.1 (Van de Peer and De Wachter, 1997). The bar indicates 10% estimated sequence divergence. Bacillus subtilis was used as outgroup (not shown). The width of the triangles is proportional to the number of genera within each cluster. The purple color indicates clusters where phototrophic bacteria occur: purple sulfur photosynthetics (Chromatiaceae and Ectothiorhodospiraceae) in the “Gammaproteobacteria,” purple nonsulfur photosynthetics and aerobic phototrophs in the “Alphaproteobacteria” and “Betaproteobacteria.” Shaded purple triangles indicate that most representatives of phototrophic “Alphaproteobacteria” and “Betaproteobacteria” are phylogenetically related to nonphototrophic, strictly chemotrophic bacteria belonging to the respective families or groups. Aerobic phototrophic Proteobacteria were reported among the Acetobacteraceae (sensu lato), the Sphingomonadaceae, the Rhodobacter, Methylobacterium and Bradyrhizobium groups (all “Alphaproteobacteria”), and in the genus Roseateles (“Betaproteobacteria,” family Comamonadaceae).

Table 5. Some Proteobacteria involved in human and animal disease. Familya

Disease (site of action)

Risk groupb 2 2 3

Ehrlichiaceae Rickettsiaceae Rickettsiaceae Rickettsiaceae

Cat-scratch disease, and bacillary angiomatosis Trench fever Abortion (genital tract of animals), and brucellosis in man Human ehrlichiosis Scrub typhus Rocky Mountain spotted fever Typhus fever

Alcaligenaceae “Burkholderiaceae” “Burkholderiaceae” Neisseriaceae Neisseriaceae Alcaligenaceae

Whooping cough (respiratory tract) Glander disease in equines Melioidosis Gonorrhoea (genital tract) Meningitis (central nervous system) Endometritis in mares

2 3 3 2 2 2

“Coxiellaceae” Enterobacteriaceae

Q-fever

3 2 2 3

Enterobacteriaceae Enterobacteriaceae Vibrionaceae Enterobacteriaceae

Diarrhea Urinary tract infections e.g., O157:H7, causing hemorrhagic diarrhea and kidney failure Tularemia (skin and lymph nodes) Meningitis, pericarditis, and pneumonia Legionnaires’ disease (respiratory tract) Nosocomial infections (skin and respiratory tract, urinary tract) Typhoid fever (gastrointestinal tract) Dysenteria (gastrointestinal tract) Cholera (gastrointestinal tract) Plague (blood)

Campylobacteraceae

Diarrhea

“Helicobacteraceae”

Duodenal and gastric ulcers, and gastric carcinoma

Proteobacterial class and species “Alphaproteobacteria” Bartonella henselae Bartonella quintata Brucella melitensis Ehrlichia chaffeensis Orientia tsutsugamushi Rickettsia rickettsii Rickettsia prowazekii “Betaproteobacteria” Bordetella pertussis Burkholderia mallei Burkholderia pseudomallei Neisseria gonorrhoeae Neisseria meningitidis Taylorella equigenitalis “Gammaproteobacteria” Coxiella burnetii Escherichia coli intestinal variants uropathogenic variants verocytotoxigenic strains Francisella tularensis Haemophilus influenzae Legionella pneumophila Pseudomonas aeruginosa Salmonella typhi Shigella dysenteriae Vibrio cholerae Yersinia pestis “Epsilonproteobacteria” Campylobacter coli, C. jejuni Helicobacter pylori

Bartonellaceae Bartonellaceae Brucellaceae

“Francisellaceae” Pasteurellaceae Legionellaceae Pseudomonadaceae

2 3 3 3

3 2 2 2 3 3 2 3 2 2 2

a

According to Bergey’s Manual of Systematic Bacteriology (Garrity and Holt, 2001). See also Fig. 1. Quotation marks are used for names which have not yet been validated (as of mid 2002). b According to EU-directive 2000/54/EG; see also (http://www.dsmz.de/).

Table 6. Some selected plant diseases caused by Proteobacteria. Proteobacterial class and species “Alphaproteobacteria” Agrobacterium rhizogenes Agrobacterium tumefaciens “Candidatus Liberibacter asiaticus” “Betaproteobacteria” Acidovorax anthurii Burkholderia cepacia Burkholderia glumae Ralstonia solanacearum Xylophilus ampelinus “Gammaproteobacteria” Brenneria (Erwinia) salicis Brenneria nigrifluens Erwinia amylovora Erwinia stewartii Pectobacterium (Erwinia) carotovorum Pseudomonas agarici Pseudomonas marginalis Pseudomonas savastanoi Pseudomonas syringae

Familya

Disease (symptoms)

Rhizobiaceae Rhizobiaceae in cluster of Rhizobiaceae, Bartonellaceae, etc.

Hairy root Crown gall Greening disease on citrus (a phloem-restricted disease)

Comamonadaceae “Burkholderiaceae” “Burkholderiaceae” “Ralstoniaceae” Comamonadaceae

Leaf-spot on Anthurium Soft rot (sour skin on onion) Sheath necrosis on rice Moko disease on banana (vascular wilt) Necrosis and canker on grapevine

Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Enterobacteriaceae Pseudomonadaceae Pseudomonadaceae Pseudomonadaceae Pseudomonadaceae

Watermark disease on willow Bark canker on Persian walnut (Juglans regia) Fire blight on pome fruit (vascular wilt) Stewart’s wilt on corn (vascular wilt) Soft rot Spots on mushrooms Soft rot (pink eye) on potato Galls on olive trees Wildfire on tobacco, haloblight on beans, spots on tomato and pepper (blights and spots)

Table 6. Continued Proteobacterial class and species Pseudomonas syringae Xanthomonas campestris Xanthomonas citri Xanthomonas oryzae Xanthomonas populi Xanthomonas translucens Xanthomonas vesicatoria Xylella fastidiosa

Familya Pseudomonadaceae “Xanthomonadaceae” “Xanthomonadaceae” “Xanthomonadaceae” “Xanthomonadaceae” “Xanthomonadaceae” “Xanthomonadaceae” “Xanthomonadaceae”

Disease (symptoms) Canker on stone fruit Black rot on crucifers (vascular wilt) Canker on citrus Blight on rice Canker on poplar trees Blight on cereals Spots on tomato and pepper Pierce’s disease (e.g., on grapevine)

a

According to Bergey’s Manual of Systematic Bacteriology (Garrity and Holt, 2001). See also Fig. 1. Quotation marks are used for names which have not yet been validated (as of mid 2002).

Table 7. Some Proteobacteria involved in industrial and biotechnological processes. Proteobacterial class, genus or species “Alphaproteobacteria” Acetobacter aceti Acetobacter xylinus Agrobacterium Gluconobacter oxydans Rhizobium Rhodobacter capsulatus Zymomonas mobilis “Betaproteobacteria” Ralstonia eutropha “Gammaproteobacteria” Azotobacter

Familya Acetobacteraceae Acetobacteraceae Rhizobiaceae Acetobacteraceae Rhizobiaceae “Rhodobacteraceae” Sphingomonadaceae

Industrial product or process Vinegar Cellulose membranes Plant engineering (Ti-plasmid) Oxidation of sorbitol (for vitamin C production) Inoculants for nodule formation on leguminous plants (N2-fixation) Production of hydrogen gas Ethanol

“Ralstoniaceae”

Poly-β-hydroxybutyrate (bioplastics) and singlecell protein

Pseudomonadaceae

Alginates (polysaccharide) and poly-βhydroxybutyrate (bioplastics) Production of hydrogen gas Biological control of frost damage Production of heterologous proteins (e.g., insulin, interferon, and antiviral vaccines) Luciferase (lux-genes) Oxidation of aliphatic and aromatic compounds Active metal mining (bioleaching) Xanthan (polysaccharide)

Chromatium Erwinia herbicola Escherichia coli

Chromatiaceae Enterobacteriaceae Enterobacteriaceae

Photobacterium Pseudomonas Acidithiobacillus ferrooxidans Xanthomonas campestris

Vibrionaceae Pseudomonadaceae Acidithiobacillus group “Xanthomonadaceae”

a

According to Bergey’s Manual of Systematic Bacteriology (Garrity and Holt, 2001). See also Fig. 1. Quotation marks are used for names which have not yet been validated (as of mid 2002).

Table 8. The “Alphaproteobacteria”: orders, families and number of genera.a Order Rhodospirillales Rickettsiales

“Rhodobacterales” “Sphingomonadales” Caulobacterales “Rhizobiales”

a

Family

Number of genera

Rhodospirillaceae Acetobacteraceae Rickettsiaceae Ehrlichiaceae “Holosporaceae” “Rhodobacteraceae” Sphingomonadaceae Caulobacteraceae Rhizobiaceae Bartonellaceae Brucellaceae “Phyllobacteriaceae” “Methylocystaceae” “Beijerinckiaceae” “Bradyrhizobiaceae” Hyphomicrobiaceae “Methylobacteriaceae” “Rhodobiaceae”

10 12 3 5 7 20 9 4 7 1 3 6 3 3 8 19 3 1

According to the second edition of Bergey’s Manual of Systematic Bacteriology (Garrity and Holt, 2001). Quotation marks are used for names which have not yet been validated (as of mid 2002).

Introduction to the Proteobacteria

BChl in some members of the “Alphaproteobacteria” (e.g., Erythrobacter and Roseobacter) is an atavistic trait that remained functional after the aerobic bacteria evolved from their anaerobic phototrophic ancestors (Stackebrandt et al., 1996; Yurkov, 2001). Thus, aerobic phototrophic bacteria could represent an intermediate phase of evolution from anaerobic purple phototrophs to nonphotosynthetic aerobic chemotrophs. During the last decade, the classification of a number of classical phototrophic genera, such as Rhodospirillum and Rhodopseudomonas, underwent considerable changes, which are in agreement with 16S rDNA sequence data, morphological parameters, internal membrane structures as well as important chemotaxonomic parameters, such as the composition of cellular fatty acids, ubiquinones and cytochrome c-type structures. Most of the photosynthetic PNS bacteria are also capable of nitrogen fixation. Classical chemoorganotrophs (such as Sphingomonas), as well as typical acidophiles (e.g., Acetobacter) and methylotrophs (e.g., Methylobacterium) belong to the “Alphaproteobacteria.” A great number of α-class members live in association with eukaryotes: some are indeed pathogenic for humans and animals (e.g., Brucella) or plants (e.g., Agrobacterium), and others display an obligate parasitic lifestyle, cause diseases in humans and mammals, are transmitted by insect or tick bites (e.g., the Rickettsiaceae), or live symbiotically in the roots of leguminous plants (e.g., Rhizobium and Bradyrhizobium) and play a key role in atmospheric nitrogen fixation. Hence, these “Alphaproteobacteria” thrive in widely divergent habitats and exert a significant impact in the biosphere of our planet. The evolutionary roots of the mitochondrion are within the α-class (see Symbiotic, Parasitic and Not-Yet Cultured Proteobacteria, and the Alphaproteobacterial Origin of Mitochondria), and the understanding of the molecular aspects of plant tumor induction by Agrobacterium tumefaciens led to revolutionary applications in plant agriculture, where plant genetic engineers have used the natural transformation system of Agrobacterium as a vector for the introduction of foreign DNA into plants (Birch, 1997). The Acetobacteraceae and the Rhodospirillaceae appear to be the deeper branching lineages among the “Alphaproteobacteria” (Ludwig and Klenk, 2001). In our analysis (Fig. 1), the Ehrlichiaceae and the Rickettsiaceae form also one of the deeper branches. Acetobacteraceae and Related Groups The Acetobacteraceae form a clearly separate lineage of acidophilic bacteria encompassing the classical vinegar-producing Acetobacter, together with Gluconobacter, Gluconaceto-

13

bacter, Acidimonas, Acidiphilium, Asaia and some other genera, as well as the phototrophic PNS bacterium Rhodopila. Some of the Gluconacetobacter species are able to fix nitrogen and live in association with sugar cane (G. diazotrophicus) or coffee plants (G. azotocaptans; Fuentes-Ramirez et al., 2001). Rickettsiaceae, Ehrlichiaceae and the Holospora Group The Rickettsiaceae, the Anaplasmataceae and the Ehrlichiaceae form a distinct lineage among the “Alphaproteobacteria” and consist of small intracellular parasitic bacteria, such as Rickettsia (causing typhus and Rocky Mountain spotted fever), Ehrlichia (causing human granulocytic ehrlichiosis and Potomac fever in horses), Anaplasma (infecting erythrocytes of ruminants), and the remarkable Wolbachia, involved in parthenogenesis in various arthropods. Although these bacteria can infect various vertebrate hosts, their vectors and reservoirs are predominantly ticks and trematodes, except for the wolbachiae, which are highly promiscuous for diverse invertebrate hosts and are also found in a variety of helminths. On the basis of 16S rDNA gene and groESL operon sequence results, Dumler et al. (2001) proposed a reorganization of the genera into the families Rickettsiaceae and Anaplasmataceae: members of the Rickettsiaceae grow in the cytoplasm or nucleus of their eukaryotic host cells and the family is restricted to the genera Rickettsia and Orientia, whereas members of the Anaplasmataceae replicate while enclosed in a eukaryotic host cell membrane-derived vacuole, and the family was broadened to include all the alphaproteobacterial species of the genera Ehrlichia, Anaplasma, Cowdria, Wolbachia and Neorickettsia (Dumler et al., 2001). Various endosymbiotic bacteria of protozoa, such as Paramecium, were allocated to the genera Holospora, Caedibacter and Lyticum forming the Holospora rDNA lineage, which is distantly related to the group formed by the rickettsiae (Fig. 1). The evolutionary roots of the eukaryotic mitochondrion are among the rickettsiae (Symbiotic, Parasitic and Not-Yet Cultured Proteobacteria, and the Alphaproteobacterial Origin of Mitochondria). Rhodospirillaceae The Rhodospirillaceae lineage corresponds to the α-1 subgroup of Woese et al. (1984a) and encompasses at least seven genera of spiral-shaped phototrophic PNS bacteria (e.g., Rhodospirillum, Phaeospirillum, Rhodospira and Rhodovibrio). Previously, these genera (except Rhodospira) were species assigned to the broad and heterogeneous genus Rhodospirillum (Imhoff et al., 1998). The phototrophic rhodospirillae practice a photoorganotrophic type of metabolism, where a simple fatty

14

K. Kersters et al.

acid or amino acid is the carbon source and light the energy source. Most of the phototrophic rhodospirillae can also fix nitrogen gas, a property that is also characteristic for members of the genus Azospirillum, belonging to the Rhodospirillaceae rDNA-lineage. The azospirillae are nonphototrophic N2-fixing bacteria, widely distributed in the rhizosphere of tropical and subtropical grasses where they seem to enhance the growth of these plants. Magnetotactic bacteria belonging to the genus Magnetospirillum are also members of this phylogenetic lineage, together with some former Aquaspirillum species. Sphingomonadaceae Proteobacteria containing glycosphingolipids in their cell envelopes belong to the phylogenetic lineage of the sphingomonads, forming a versatile group of aerobic bacteria occurring in various environments such as soil, water and clinical specimens. This lineage corresponds to the α-4 subgroup. The genus Sphingomonas contained at least 20 species and was recently split on the basis of phylogenetic and chemotaxonomic analyses into four different genera (Sphingomonas sensu stricto, Sphingobium, Novosphingobium and Sphingopyxis; Takeuchi et al., 2001). Some of the species belonging to these genera can metabolize various aromatic compounds and possess promising biotechnological properties. The Sphingomonadaceae comprise also the genera Erythrobacter, Erythromicrobium and Porphyrobacter, which are aerobic chemoorganotrophs, although their cells contain bacteriochlorophyll a and carotenoids. The genus Zymomonas is related to the sphingomonads. Zymomonas strains occur in palm sap and pulque, where they ferment sugars to high concentrations of ethanol by using the Entner-Doudoroff pathway, leading to the alcoholic beverages palm wine and tequila, respectively (Swings and De Ley, 1977; Sahm et al., 2001). Rhodobacter Group The Rhodobacter group is a heterogeneous phylogenetic lineage among the “Alphaproteobacteria,” including a number of photosynthetic rod-shaped PNS bacteria as well as numerous chemoorganotrophs. It corresponds to the α-3 group (Woese et al., 1984a). Typical photosynthetic PNS bacteria of this lineage belong to the genera Rhodobacter and Rhodovulum, containing freshwater and marine species, respectively. Rhodobacter sphaeroides and R. capsulatus were extensively used for genetic studies of bacterial photosynthesis. The facultative chemolithotroph Paracoccus denitrificans belongs also here; it is a remarkable and versatile bacterium, being able to grow chemolithoautotrophically at the expense of hydrogen gas or reduced inorganic sulfur compounds as electron

donors, carbon dioxide and oxygen, but it can also grow chemoorganotrophically with various organic compounds as sole carbon source. Some hyphal, budding and prosthecate aerobic bacteria such as Hyphomonas, Hirschia and Gemmobacter belong to this lineage, as well as the budding Staleya and Antarctobacter, which were isolated from a hypersaline, heliothermal antarctic meromictic lake (Labrenz et al., 1998; Labrenz et al., 2000). Gram-negative cocci typically occurring in regular packages of tetrads and isolated from activated sludge biomass were assigned to the genus Amaricoccus, belonging to the Rhodobacter lineage (Maszenan et al., 1997). The Rhodobacter group is peripherally related to some of the most abundant cultured marine species and strains from some terrestrial halophilic lake environments. These organisms, frequently referred to as the Roseobacter group, include in addition to the aerobic bacteriochlorophyllcontaining genera (such as Roseobacter, Roseovarius and Rubrimonas) the genera Antarctobacter, Ketogulonicigenium, Methylarcula, Octadecabacter, Sagitulla, Silicibacter, and Sulfitobacter, as well as the phylogenetically heterogeneous genus Ruegeria. Caulobacteraceae The Caulobacteraceae are aquatic chemoorganotrophic and aerobic bacteria often attaching to surfaces with a stalk at one end and forming polarly flagellated swarming cells at the other end. Stalks of several cells may form rosettes. Caulobacters occur typically in freshwater and marine habitats with low nutrient levels. The rod-shaped Brevundimonas strains, which were previously allocated to the genus Pseudomonas, belong also to the Caulobacteraceae. Rhizobiaceae, Bartonellaceae, Brucellaceae and Phyllobacterium Group This large and complex rDNA-cluster contains at present at least 15 genera and 70 species and corresponds to the α-2 subgroup of Woese et al. (1984a), together with the adjacent lineage formed by the Hyphomicrobiaceae and the Bradyrhizobium, Methylobacterium and Methylocystis rDNA groups (Fig. 1). One common trait of bacteria belonging to this cluster is that most of them interact with eukaryotes: agrobacteria are plant pathogens causing crown-gall or hairy-root disease (tumors) on various dicotyledonous plants; the rhizobia induce nodules on roots or stems of leguminous plants and live symbiotically in these nodules where they reduce atmospheric nitrogen; and the bartonellae and brucellae are pathogenic for humans or animals. The fast growing rhizobia classified in the genera Rhizobium, Allorhizobium, Mesorhizobium and Sinorhizobium form a major rDNA cluster, together with

Introduction to the Proteobacteria

the plant pathogenic genus Agrobacterium and the genus Phyllobacterium, whose members were isolated from leaf nodules of various plants belonging to the Myrsinaceae and Rubiaceae. The genus Mesorhizobium (more related to Phyllobacterium) and members of the genus Sinorhizobium form also a distinct clade among the rhizobia. According to 16S rDNA analysis, various Rhizobium species as well as Allorhizobium are highly related to the plant pathogenic agrobacteria. Recently, Young et al. (2001) proposed inclusion of all the species of Agrobacterium and Allorhizobium in the genus Rhizobium. The rRNA-based classification of the rhizobia is generally supported by sequence analysis of atpD and recA genes (Gaunt et al., 2001). The genes for nodule formation (nod genes), nitrogen fixation (nif genes) and tumor induction (in the case of agrobacteria) are localized on large plasmids. Species of the genus Bartonella occur in the blood of man and mammals; they are often vector borne, but can also be transmitted by animal scratches or bites. Bartonellae are considered to be emerging human pathogens. The type species Bartonella bacilliformis is a fastidious hemophilic organism that invades and destroys human red blood cells and is transmitted by a sandfly. Bartonella quintana (Brenner et al., 1993) was previously classified in the genus Rochalimaea and causes trench fever, a disease transmitted by lice and afflicting, e.g., soldiers during the First World War (1914–1918). Bartonella henselae (formerly Rochalimaea henselae) is nowadays recognized as the causative agent of cat scratch disease (Table 5). Brucellae develop intracellularly and cause worldwide infections (brucellosis) in humans and a great number of animals such as cattle, pigs, dogs, and even marine mammals. Hyphomicrobiaceae, the Bradyrhizobium, Methylobacterium, Methylocystis and Related Groups The family Hyphomicrobiaceae contains hyphal, prosthecate and budding bacteria; most of them (e.g., Hyphomicrobium and Pedomicrobium) are chemoorganotrophs and prefer to grow on one-carbon compounds such as methanol, whereas others are phototrophic (e.g., Rhodomicrobium). Hyphomicrobium is well adapted to grow in oligotrophic freshwater habitats. Other members of the family are Xanthobacter, a versatile soil bacterium capable of nitrogen fixation and autotrophic growth in an atmosphere of hydrogen, oxygen and carbon dioxide, and Azorhizobium caulinodans, a nitrogen-fixing bacterium living symbiotically in the stem nodules of some leguminous plants such as Sesbania. A large subcluster within this complex lineage is formed by the Bradyrhizobium group, includ-

15

ing the slow-growing rhizobia nodulating soy bean and other leguminous plants and the stem-nodulating phototrophic bradyrhizobia (Molouba et al., 1999; Giraud et al., 2002), the photosynthetic PNS Rhodopseudomonas, the ecologically important chemolithotrophic nitrobacters oxidizing nitrite to nitrate, as well as the opportunistic Afipia species. The Methylobacterium and Methylocystis groups contain various methanotrophic and methylotrophic bacteria, mostly utilizing the serine pathway for the assimilation of one-carbon intermediates. Some of these methanotrophs may have biotechnological applications because they can utilize chloromethanes from polluted environments, whereas others such as Methylocapsa are acidophilic and can fix atmospheric nitrogen (Dedysh et al., 2002). A subgroup of the methylotrophic methylobacteria has been shown to nodulate Crotolaria legumes and fix nitrogen (Sy et al., 2001). Other members of this rDNA lineage are the genera Beijerinckia, consisting of aerobic, acid-tolerant free-living nitrogen-fixing rods occurring mainly in tropical acidic soils, and Rhodobium, which contains marine budding phototrophic PNS bacteria.

The “Betaproteobacteria” The “Betaproteobacteria” (at least 75 genera and 220 species) clearly represent a monophyletic group within the larger phylogenetic lineage composed of the β-γ proteobacterial complex, named Chromatibacteria by Cavalier-Smith (Cavalier-Smith, 2002; Table 2). From the metabolic, morphological and ecological viewpoint, the “Betaproteobacteria” are very heterogeneous. They contain some purple nonsulfur (PNS) phototrophs (Fig. 1, purple triangles), various chemolithotrophs, some methylotrophs, a great number of chemoorganotrophs, some nitrogen-fixing bacteria, and some important plant-, human- and animal pathogens (see Tables 5 and 6). Their morphologies can vary from rods or cocci to spiral and sheathed cells. Some members of the “Betaproteobacteria” are of biotechnological interest owing to their biodegradation properties. Recently nitrogen-fixing “Betaproteobacteria” were described that nodulate the roots of legumes (Chen et al., 2001; Moulin et al., 2001). More detailed information concerning the group can be found in the chapters dealing with the individual families and genera of the “Betaproteobacteria.” Table 9 gives an overview of the major phylogenetic groups (see also Fig. 1) and the names of the six orders and 12 families of the “Betaproteobacteria” as they are listed in the 2nd edition of Bergey’s Manual of Systematic Bacteriology (Garrity, 2001a).

16

K. Kersters et al.

Table 9. The “Betaproteobacteria”: orders, families and number of genera.a Order “Burkholderiales”

“Hydrogenophilales” “Methylophilales” “Neisseriales” “Nitrosomonadales”

“Rhodocyclales”

Family “Burkholderiaceae” “Ralstoniaceae” “Oxalobacteraceae” Alcaligenaceae Comamonadaceae “Hydrogenophilaceae” “Methylophilaceae” Neisseriaceae “Nitrosomonadaceae” Spirillaceae Gallionellaceae “Rhodocyclaceae”

Number of genera 4 1 5 6 15 2 3 14 2 1 1 6

a According to the second edition of Bergey’s Manual of Systematic Bacteriology (Garrity and Holt, 2001). Quotation marks are used for names which have not yet been validated (as of mid 2002).

Although the majority of the phototrophic PNS bacteria belong to the “Alphaproteobacteria,” some PNS species are members of the Rhodocyclus group or the family Comamonadaceae of the “Betaproteobacteria.” Similarly to the previously mentioned PNS “Alphaproteobacteria,” the PNS betaproteobacterial phototrophs are phylogenetically intermingled with nonphototrophs (Fig. 1, visualized by purple shaded areas) and can be clearly differentiated from the PNS “Alphaproteobacteria” (e.g., Rhodobacter, Rhodobium, etc.) on the basis of fatty acid and quinone composition as well as cytochrome c sequences (Imhoff, 2001a). Alcaligenaceae, Comamonadaceae, Burkholderia, Oxalobacter and Related Groups The Alcaligenaceae, Comamonadaceae, and the Burkholderia-, Ralstonia- and Oxalobactergroups are phylogenetically more closely related to each other than to other 16S rDNA branches of the “Betaproteobacteria” (Fig. 1). A number of clinically important taxa belong to this lineage: members of the genus Alcaligenes are mostly saprophytic, but behave often as nosocomial bacteria, whereas the closely related Bordetella pertussis (Table 5) causes whooping cough in humans (other Bordetella species are pathogenic for animals). Taylorella is responsible for endometritis in mares, Pelistega is associated with a respiratory disease in pigeons (Vandamme et al., 1998), and Brackiella oedipodis (Willems et al., 2002) was recently reported as the causal agent of endocarditis in a small neotropical primate. The former “acidovorans” and “solanacearum” rRNA groups of the genus Pseudomonas sensu lato (i.e., respectively, rRNA groups III and II of Palleroni [1984]) belong to the “Beta-

proteobacteria.” The “acidovorans” group has at present the status of the family Comamonadaceae, a phylogenetically coherent but physiologically heterogeneous group of prokaryotes encompassing genera such as Acidovorax, Comamonas, Delftia, Hydrogenophaga, Variovorax, Xylophilus, Rhodoferax, Roseateles, Rubrivivax, Leptothrix and Sphaerotilus (Willems et al., 1991a; Willems et al., 1991b). The latter two genera are heterotrophic sheathed bacteria, which may appear yellow or dark brown owing to the deposition of iron and manganese oxides, whereas Rhodoferax and Rubrivivax are typical PNS photosynthetic bacteria. Some Comamonadaceae, such as Acidovorax facilis, Hydrogenophaga flava and Variovorax paradoxus, are able to grow chemolithotrophically at the expense of hydrogen gas oxidation. Xylophilus ampelinus causes necrosis and canker on grapevines. Other representatives, e.g., belonging to Comamonas or Delftia, can degrade aromatic compounds (via the meta-cleavage of the aromatic ring) and are important in the biodegradation of toxic wastes. Roseateles was described as the first obligate aerobic betaproteobacterium containing bacteriochlorophyll a (Suyama et al., 1999). However, light does not support growth of Roseateles strains under anaerobic conditions and in this sense Roseateles resembles physiologically the aerobic bacteriochlorophyll a-containing “Alphaproteobacteria,” such as Erythrobacter and Erythromicrobium (Sphingomonadaceae). The former “Pseudomonas solanacearum group” is composed of the genera Burkholderia, Ralstonia and a few others. Some of these bacteria are typical plant pathogens (e.g., Ralstonia solanacearum, causing wilt on many cultivated plants; Table 6), whereas others are important animal and human pathogens. Burkholderia mallei and B. pseudomallei have been classified as risk group 3 organisms (Table 5) because they cause respectively glanders disease in horses and melioidosis, a disease endemic in animals and humans in Southeast Asia. Burkholderia cepacia seems ubiquitous and is both friend and foe to humans (Govan et al., 2000): it is associated with plants (the original isolate was reported by Burkholder [1950] as the causative agent of bacterial rot of onion bulbs), occurs in soil and water, protects crops from bacterial and fungal infections, and is also frequently reported in the environment of patients, particularly cystic fibrosis patients, where B. cepacia infections have a considerable impact on their morbidity and mortality, as well as on their social life. The remarkable genomic heterogeneity among the B. cepacia strains isolated from various ecological niches makes their correct identification problematic (Coenye et al., 2001). It was demonstrated that presumed B. cepacia strains isolated

Introduction to the Proteobacteria

from cystic fibrosis patients and other sources belong to at least nine distinct genomic species or genomovars (Vandamme et al., 1997; Coenye et al., 2001). At least two of these genomovars are often implicated in the transmission of B. cepacia between cystic fibrosis patients (Mahenthiralingam et al., 2000; LiPuma et al., 2001). The extreme metabolic versatilty of B. cepacia could be applied in the bioremediation of recalcitrant xenobiotics (Parke and GurianSherman, 2001). However, more caution is undoubtedly needed in this area, because many strains used or under development for biocontrol or bioremediation purposes are taxonomically poorly characterized, causing a potential hazard to the cystic fibrosis community worldwide (Parke and Gurian-Sherman, 2001). The related genus Ralstonia is also unusual as it harbors important plant pathogens, opportunistic human pathogens, as well as organisms of considerable biotechnological interest because of their potential for biodegradation of xenobiotics and recalcitrant compounds. Moreover, Chen et al. (2001) described Ralstonia taiwanensis as a betaproteobacterium capable of root nodule formation and nitrogen fixation in a leguminous plant (Mimosa). Together with the Burkholderia-like strains described by Moulin et al. (2001), these are the first betaproteobacteria known to be capable of root nodule formation and nitrogen fixation. Ralstonia eutropha (previously classified as Alcaligenes eutrophus) is a well-known and intensely studied facultative chemolithotroph (a typical “Knallgas” bacterium) that can grow at the expense of hydrogen gas, CO2 and O2 (Aragno and Schlegel, 1992). When grown under appropriate conditions, R. eutropha is also an excellent producer of the bioplastic poly-βhydroxybutyrate and similar polyhydroxyalkanoates (Table 7). Some toxic-metal-resistant bacteria isolated from industrial biotopes were recently allocated to the genus Ralstonia (Goris et al., 2001). The latter organisms can be exploited to recycle soils polluted by toxic metals, such as mercury, cadmium or nickel. The Oxalobacter group is composed of Herbaspirillum (a diazotroph colonizing endophytically roots, stems or leaves of graminaceous plants), Janthinobacterium (one of the known violacein-producing organisms), Oxalobacter and a few other genera. Oxalobacter formigenes occurs in the rumen or the large bowel of humans and animals; it performs a unique fermentation of oxalate to formate and CO2, whereby ATP is generated via a membranelinked proton-translocating ATPase. Hydrogenophilus Group The genus Hydrogenophilus harbors thermophilic (optimum growth temperature 50–60°C), facultatively

17

chemolithoautotrophic hydrogen-oxidizing rods that form a distinct lineage among the “Betaproteobacteria” (Hayashi et al., 1999). The classification of the colorless chemolithotrophic S-oxidizing Thiobacillus species has been thoroughly revised by Kelly and Wood (2000). The type species T. thioparus together with a few other Thiobacillus species belongs to the “Betaproteobacteria” and is distantly related to the Hydrogenophilus group. The majority of the other Thiobacillus species belong to the “Gammaproteobacteria” and were reassigned to three other genera (see The “Gammaproteobacteria”). Because they are able to leach metals from ore, Thiobacilli (e.g., Acidithiobacillus species) are used in processing low-grade metal ores. Methylophilus, Nitrosomonas and Related Groups The majority of the one-carbon utilizers (methanotrophs) belong either to the “Alphaproteobacteria” or to the “Gammaproteobacteria.” However, three methylotrophic genera (Methylobacillus, Methylophilus and Methylovorus) group in the Methylophilus rRNA lineage of the “Betaproteobacteria.” These are phylogenetically distantly related to the Nitrosomonas lineage. The aerobic chemolithoautotrophic ammonia-oxidizing bacteria such as the rod-shaped Nitrosomonas and the spiral-shaped Nitrosospira constitute a monophyletic lineage among the “Betaproteobacteria.” Spirillum volutans is distantly related to this group, as well as the Fe2+-oxidizing chemolithotrophic Gallionella ferruginea, already known since 1836 for its peculiar bean-shaped cells forming very long spiral stalks composed of ferric hydroxide (Ehrenberg, 1836). Rhodocyclus Group The Rhodocyclus lineage harbors besides PNS bacteria (e.g., Rhodocyclus purpureus) also diazotrophic rhizosphere bacteria such as Azoarcus and Azospira. Several strains of Azoarcus and the related Thauera can degrade aromatic compounds including chlorobenzoates. Thauera selenatis is capable of nitrate and selenate respiration and another member of this group, Zoogloea ramigera (Rossellomora et al., 1993), is considered to be a typical floc-forming prokaryote during the activated sludge process in sewage treatment plants (Juretschko et al., 2002). Also Quadricoccus (a large coccus occurring in tetrads; Maszenan et al., 2002) is found in the activated sludge biomass where it synthesizes polyphosphate and polyhydroxyalkanoates. Dechloromonas and Dechlorosoma are newly described genera of perchlorate-reducing bacteria, associated, e.g., with the manufacture and dismantling of ammunition (Achenbach et al., 2001).

18

K. Kersters et al.

Neisseriaceae The most prominent human pathogens of the “Betaproteobacteria” belong to the genus Neisseria, containing the two wellknown species N. gonorrhoeae and N. meningitidis. Strains of these organisms have been extensively studied by multi-locus-sequencing-typing (MLST; Achtman, 1998; Maiden et al., 1998). Other clinical bacteria, such as Kingella and Eikenella, belong also to this phylogenetic group together with Chromobacterium, which produces the purple pigment violacein. The mammalian oral commensals Alysiella and Simonsiella belong also to Neisseriaceae and are unique because their filament-forming cells show a dorsal-ventral asymmetry and display a gliding motility on the epithelial cells in the oral cavity and upper respiratory tract of their host. These bacteria might have coevolved with their mammalian hosts during the past 100 million years (Hedlund and Staley, 2002).

The “Gammaproteobacteria” Most 16S rDNA trees show that the members of the “Gammaproteobacteria” represent a monophyletic group which includes in fact also the “Betaproteobacteria” as a major line of descent (see Fig. 1). The “Gammaproteobacteria” form the largest proteobacterial group (at least 180 genera and 750 species), including the phytopathogenic Xanthomonas group as a border-line member. Depending upon the rDNA-treeing method used, the Xanthomonas rDNA-group appears peripherally linked to either the β- or the γ-class and can be considered as a sister group of the “Betaproteobacteria” (Ludwig and Klenk, 2001). The major phylogenetic groups (Fig. 1) and the names of the 13 orders and 20 families of the “Gammaproteobacteria” as listed in the 2nd edition of Bergey’s Manual of Systematic Bacteriology (Garrity, 2001a) are given in Table 10. The individual chapters on the families and genera of this class can be consulted for more detailed information. The “Gammaproteobacteria” contain the photosynthetic purple sulfur (PS) bacteria (Chromatiaceae and Ectothiorhodospiraceae; Table 10; Fig. 1) together with a great number of familiar chemoorganotrophic bacterial groups, such as the Enterobacteriaceae, Legionellaceae, Pasteurellaceae, Pseudomonadaceae, Vibrionaceae, and also some chemolithotrophic mostly sulfuror iron-oxidizing prokaryotes. The class harbors some important human and animal pathogens (Table 5). It is noteworthy that the family Enterobacteriaceae is known since 1937 as a classical phenotypic group (Rahn, 1937), which remains fully supported by modern molecular taxonomy. On the other hand, the tradional group of the pseudomonads turned out to be phylogenetically

Table 10. The “Gammaproteobacteria”: orders, families and number of genera.a Order “Chromatiales” “Xanthomonadales” “Cardiobacteriales” “Thiotrichales”

“Legionellales” “Methylococcales” “Oceanospirillales” Pseudomonadales “Alteromonadales” “Vibrionales” “Aeromonadales” “Enterobacteriales” “Pasteurellales”

Family

Number of genera

Chromatiaceae Ectothiorhodospiraceae “Xanthomonadaceae” Cardiobacteriaceae “Thiotrichaceae” “Piscirickettsiaceae” “Francisellaceae” Legionellaceae “Coxiellaceae” Methylococcaceae “Oceanospirillaceae” Halomonadaceae Pseudomonadaceae Moraxellaceae Alteromonadaceae Vibrionaceae Aeromonadaceae Succinivibrionaceae Enterobacteriaceae Pasteurellaceae

22 5 8 3 9 5 1 1 2 6 6 6 15 3 11 6 2 4 41 6

a According to the second edition of Bergey’s Manual of Systematic Bacteriology (Garrity and Holt, 2001). Quotation marks are used for names which have not yet been validated (as of mid 2002).

extremely heterogeneous, because its members are scattered over the “Alphaproteobacteria,” “Betaproteobacteria” and “Gammaproteobacteria.” The genus Pseudomonas is at present restricted to all species phylogenetically related to its type species Pseudomonas aeruginosa, a member of the “Gammaproteobacteria,” whereas all the other pseudomonads belonging to the α- and β-classes have been allocated to new genera such as Brevundimonas, Sphingomonas, Comamonas, Burkholderia, Ralstonia, etc. (see The “Alphaproteobacteria,” The “Betaproteobacteria”). An analogous taxonomic history concerns the genus Thiobacillus, whose members are chemolithotrophs oxidizing various inorganic sulfur-compounds. Its classification was recently clarified by Kelly and Wood (2000): thiobacilli related to the type species T. thioparus belong to the “Betaproteobacteria,” whereas the Thiobacillus species belonging to the “Gammaproteobacteria” were reclassified in the genera Acidithiobacillus, Halothiobacillus and Thermithiobacillus. Chromatiaceae and Ectothiorhodospiraceae The distribution of the anoxygenic photosynthetic purple sulfur (PS) bacteria is at present restricted to two clearly distinct phylogenetic lineages of the “Gammaproteobacteria” (Fig. 1): the Chromatiaceae (25 genera), which accumulate elemental sulfur globules inside their cells,

Introduction to the Proteobacteria

and the Ectothiorhodospiraceae (7 genera), which deposit sulfur outside their cells. These prokaryotes are mainly found in illuminated anoxic zones of lakes (particularly meromictic lakes) where H2S accumulates and also in “sulfur springs,” where biologically or geochemically produced H2S can trigger the formation of massive blooms of purple sulfur bacteria (Imhoff, 2001a). All PS bacteria can utilize H2S as an electron donor for CO2 reduction, but many are also able to use other reduced sulfur compounds (e.g., thiosulfate) as photosynthetic electron donors. Ectothiorhodospiraceae are haloalkaliphilic sulfur bacteria possessing lamellar intracellular photosynthetic membrane structures, whereas most Chromatiaceae possess a vesicular type of intracellular membrane. Some nonphototrophic and alkaliphilic taxa belong also to the phylogenetic lineage of the Ectothiorhodospiraceae: for example, Nitrococcus is a halophilic nitriteoxidizing chemolithotroph, Thioalkalivibrio (Sorokin et al., 2001) contains obligately chemolithotrophic sulfur-oxidizing bacteria, and Alcalilimnicola (Yakimov et al., 2001) is a halotolerant aerobic heterotroph. Enterobacteriaceae, Aeromonadaceae, Alteromonadaceae, Pasteurellaceae, Succinivibrionaceae and Vibrionaceae The core of the γ-class is composed of the well-known enterics (see Introduction to the Family Enterobacteriaceae in the second edition), Vibrionaceae (see The Family Vibrionaceae in Volume 6), Aeromonadaceae (see The Genera Aeromonas and Plesiomonas in Volume 6), Pasteurellaceae (see The Genus Pasteurella in Volume 6) and the Alteromonadaceae (see The Genus Alteromonas and Related Proteobacteria in Volume 6), each including a large number of genera and species. Some representatives of these families are known as pathogens of humans and animals (Table 5). The Enterobacteriaceae include the well-known and thoroughly studied Escherichia coli, together with numerous inhabitants of the intestinal tract of warm-blooded animals (e.g., Salmonella and Shigella, the etiological agents of salmonellosis and shigellosis, respectively; see Table 5). Escherichia coli is an excellent indicator organism for water quality (fecal contamination). The Enterobacteriaceae comprise a relatively homogeneous phylogenetic group within this large cluster; they are mostly facultative anaerobic carbohydrate-degrading microorganisms; some perform a mixed acid fermentation, whereas others carry out the butanediol fermentation. The enterics comprise also plant pathogenic bacteria (e.g., Erwinia, Pectobacterium and Brenneria; see Table 6), the plague-causing Yersinia pestis, as well as Xenorhabdus and Photorhabdus, which are symbionts of entomopatho-

19

genic nematodes, where they live in the intestinal lumen. An interesting and somewhat distant member of the enterics is Buchnera aphidicola, a not-yet-cultured endosymbiont of aphids (Baumann et al., 1995). The symbiotic association between aphids and Buchnera seems to be obligate and mutualistic: Buchnera synthesizes tryptophan, cysteine and methionine and supplies these essential amino acids to the aphid host. A parallel evolution of Buchnera and aphids seems to have occurred, and Baumann et al. (1998) estimated the origin of this symbiotic association as 200–250 million years ago. The correlation of sequence diversity of 16S rDNA of symbionts with the age of their hosts has led to the calibration of the molecular clock of 16S rDNA in recent organisms (Moran et al., 1993), assumed to generate 1% sequence divergence within 25– 50 million years (see also Stackebrandt, 1995). Also the endosymbionts of carpenter ants (Camponotus spp.) constitute a distinct taxonomic group within the “Gammaproteobacteria” and are phylogenetically closely related to Buchnera and symbionts of tsetse flies. Comparison of the phylogenetic trees of the bacterial endosymbionts and their host species suggests a highly synchronous cospeciation process of both partners (Sauer et al., 2000). Vibrionaceae are facultative anaerobic inhabitants of brackish, estuarine and pelagic waters and sediments and form the dominant culturable microflora in the gut of molluscs, shrimps and fish. The family harbors several pathogens (e.g., Vibrio cholerae, the causal agent of cholera) as well as luminous bacteria (e.g., Photobacterium and several Vibrio species), occurring free-living in seawater, as well as being symbionts in the light organs of many fish and invertebrates (see Dunlap and Kita-Tsukamoto, 2001). Bioluminescent bacteria occur also among the genera Photorhabdus (Enterobacteriaceae) and Shewanella. The latter genus belongs to the Alteromonadaceae, which are strictly aerobic chemoorganotrophs requiring seawater for growth. Quorum-sensing plays an important role in the phenomenon of bioluminescence. Several representatives of the Pasteurellaceae are the etiological agents of various infectious diseases in humans and other vertebrates (cattle, sheep, goats and fowl). They encompass the following major genera: Pasteurella, Haemophilus, Actinobacillus and Mannheimia. Haemophilus influenzae was the first free-living organism whose entire genome (ca. 1.8 × 109 bp) was sequenced (Fleischmann et al., 1995). The Aeromonadaceae include the fish pathogen Aeromonas salmonicida. The Succinivibrionaceae (Hippe et al., 1999) are strict anaerobes fermenting glucose and other carbohydrates to succinate and acetate. They occur in the rumen of sheep and

20

K. Kersters et al.

cattle (Ruminobacter, Succinomonas and Succinivibrio) or in the feces or colon of dogs and humans (Anaerobiospirillum). Francisella and Piscirickettsia Groups The Francisella and Piscirickettsia groups form a deeply branching lineage among the “Gammaproteobacteria.” Francisella tularensis is the causal agent of tularemia, a plague-like zoonosis, spread to humans from rodents via direct contact or biting arthropods, whereas Piscirickettsia salmonis causes an epizootic disease in salmonid fishes. Members of the Piscirickettsia group display a great morphological and metabolic diversity: Thiomicrospira contains spiral-shaped chemolithotrophic sulfur-oxidizing bacteria, which are closely related to the alkaliphilic chemolithotrophic Thioalkalimicrobium (Sorokin et al., 2001). Hydrogenovibrio is a marine obligately chemolithoautotrophic hydrogen-oxidizing bacterium and Cycloclasticus harbors rod-shaped bacteria degrading aromatic hydrocarbons and occurring in marine sediments. Cardiobacteriaceae The Cardiobacteriaceae are an example of a family created (Dewhirst et al., 1990) on the basis of 16S rDNA comparisons of three taxa, whose phylogenetic relationship was at first quite unexpected. Cardiobacterium hominis is an occasional resident of the human respiratory tract and has been recovered from blood samples of patients suffering from endocarditis. Suttonella indologenes (previously classified as Kingella indologenes) has been reported in human eye infections and blood of patients with endocarditis, whereas Dichelobacter nodosus (formerly Bacteroides nodosus) is the causative agent of footrot in ruminants. Authentic kingellae belong to the “Betaproteobacteria” and authentic bacteroideae belong to the Cytophaga-Flavobacterium-Bacteroides phylum (the so-called “Bacteroidetes” according to Garrity, 2001). Moraxellaceae The Moraxellaceae encompass the genera Moraxella, Acinetobacter and Psychrobacter forming a separate lineage among the “Gammaproteobacteria” (Rossau et al., 1991). Moraxellae are commonly isolated from animals or humans and some of them are pathogenic, whereas the psychrophilic Psychrobacter species occur in the marine and antarctic environment (e.g., in antarctic ornithogenic soils; Bowman et al., 1996) and also in clinical material. The oxidase-negative acinetobacters are saprophytes occurring in soil, water and sewage, but also typically on the skin of healthy humans. Moreover, some acinetobacters can also cause nosocomial infections. Methylococcaceae The Methylococcaceae form a distinct lineage (Bowman et al., 1995)

encompassing methylotrophic bacteria such as the genera Methylobacter, Methylococcus and Methylomonas, which share the ribulose monophosphate cycle for the assimilation of reduced C1-compounds. Some of these methylotrophs form cysts as resting stages and possess internal membrane systems arranged as bundles of discshaped vesicles distributed throughout the cell, in contrast to some methylotrophs of the “Alphaproteobacteria,” whose membrane systems run along the periphery of the cells. The moderately halophilic Methylophaga belong phylogenetically to the Piscirickettsia group. Thiothrix Group The Thiothrix group contains a number of filamentous and gliding sulfuroxidizing chemolithotrophs, such as Thiothrix, Beggiatoa and Thioploca. Thiothrix forms sheathed filaments, whereas Beggiatoa forms long filaments lacking a sheath. Sulfur granules generally accumulate in the cells. The not yet axenic Thiomargarita (“sulfur pearl”) is related to Thioploca and is a real giant among the prokaryotic life forms (its spherical cells are about 500 µm wide!). Thiomargarita cells form thick mats near the coast of Namibia, where they perform an anoxic oxidation of H2S coupled to the reduction of nitrate (Schulz et al., 1999; Schulz, 2002). Leucothrix can be considered as a chemoorganotrophic counterpart of Thiothrix. Some not yet cultured sulfur-oxidizing symbiotic bacteria of bivalves are closely related to the Thiothrix group and play an important role in the nutrition of such animals, living near hydrothermal vents. Halomonadaceae The family Halomonadaceae is a large and complex group of moderately halophilic and marine bacteria encompassing the genera Halomonas, Chromohalobacter, Zymobacter and Carnimonas. Recent comparative sequence analysis of 16S and 23S rDNA of representative strains indicated (Arahal et al., 2002) that the genus Chromohalobacter forms a monophyletic group, whereas the genus Halomonas is clearly polyphyletic and needs a taxonomic re-evaluation based on a polyphasic taxonomic approach (Romanenko et al., 2002 [doi 10.1099/ijs.0.02240-0]). Pseudomonadaceae In the past (Palleroni, 1984), the family Pseudomonadaceae contained an extremely heterogeneous group of aerobic, rod-shaped and mostly polarly flagellated bacteria, phylogenetically spread over the “Alphaproteobacteria,” “Betaproteobacteria” and “Gammaproteobacteria.” Nowadays the family is restricted to some 90 Pseudomonas species around the type species Pseudomonas aeruginosa and some related genera which form a

Introduction to the Proteobacteria

separate lineage within the “Gammaproteobacteria.” The pseudomonads of the “Alphaproteobacteria” and “Betaproteobacteria” have been transferred to other genera (Willems et al., 1991a; Kersters et al., 1996; Anzai et al., 2000; Caulobacteraceae, Alcaligenaceae, Comamonadaceae, Burkholderia, Oxalobacter and Related Groups). Some authentic Pseudomonas species (e.g., P. fluorescens) produce fluorescent pigments. Most pseudomonads are versatile and can grow on a great variety of organic compounds, including aromatic hydrocarbons, because some of them possess efficient oxygenases. Such bacteria play key roles in the purification of wastewater and clean-up of oil spills. Some of the Pseudomonas species (e.g., P. syringae) are plant pathogens (Table 6), whereas P. aeruginosa and some other fluorescent pseudomonads can be involved in serious nosocomial infections (Table 5). The free-living nitrogen fixers of the genera Azotobacter and Azomonas belong also to this phylogenetic lineage, together with the cellulosedegrading Cellvibrio and a few other related genera. Oceanospirillum Group The Oceanospirillum group encompasses six genera whose members are aerobic, are moderately halophilic and generally possess curved cells. Some representatives (e.g., Neptunomonas) can degrade polycyclic aromatic hydrocarbons or long chain alkanes. The taxonomy of the genus Oceanospirillum has recently been revised (Satomi et al., 2002). Balneatrix alpica was reported as a bacterium associated with a case of pneumonia and meningitis in a spa therapy center in France (Dauga et al., 1993). Legionellaceae and Coxiella Group The clinically important legionellae occur in surface water, mud, thermally polluted lakes and streams and some (e.g., Legionella pneumophila) may enter the human respiratory tract when water is aerosolized in showers and airconditioning systems. Legionella pneumophila is a pathogen for humans causing pneumonia (Legionnaires’ disease). An obligate intracellular bacterial parasite of small free-living amoebae (previously classified as Sarcobium lyticum; Drozanski, 1991) belongs also to the genus Legionella (Hookey et al., 1996). The genera Coxiella and Rickettsiella belong to the same phylogenetic lineage as the legionellae. Although phylogenetically distinct, Coxiella and Rickettsia (the latter belonging to the “Alphaproteobacteria”) share similarities in their parasitic lifestyle. Coxiella burnetii, an obligate parasitic bacterium growing preferentially in the vacuoles of the host cells, causes Q-fever, a pneumonia-like infection, transmitted among animals by insect bites (e.g.,

21

ticks) and occasionally causes disease in humans (originally called “abattoir fever”). Rickettsiellae are widely distributed intracellular pathogens of invertebrates, including insects, crustaceans and arachnids. They have not yet been cultivated in cell-free media and grow and multiply in cell vacuoles of the fat body and hepatopancreas of their invertebrate hosts. Xanthomonas Group The plant pathogenic Xanthomonas species, Frateuria (phenotypically resembling the acetic acid bacteria of the αclass), Xylella (a phytopathogen living in the xylem of various plants) and Stenotrophomonas, together with some yellow-pigmented N2Oproducing bacteria isolated from ammoniasupplied biofilters (Finkmann et al., 2000), constitute a clearly separated phylogenetic lineage among the Chromatibacteria sensu Cavalier-Smith (2002), i.e., the large complex formed by the “Betaproteobacteria” and the “Gammaproteobacteria”; see Table 2). The ubiquitous Stenotrophomonas maltophilia is also involved in important nosocomial infections (Van Couwenberghe et al., 1997; Khan and Mehta, 2002). Depending on the treeing algorithms used and the number of rDNA-sequences included, the xanthomonads cluster peripherally linked either to the “Betaproteobacteria” or to the “Gammaproteobacteria” (Fig. 1). The first complete genome sequence published of a plant pathogenic bacterium (Simpson et al., 2000) was that of Xylella fastidiosa, a pathogen causing important diseases in citrus trees, grapevines and other plants.

The “Deltaproteobacteria” From the viewpoint of lifestyle and morphology, the δ-class is most peculiar because it contains bacteria that are typical predators on other prokaryotes (bdellovibrios), whereas other “Deltaproteobacteria” belonging to the myxobacteria display complex developmental life cycles, forming multicellular structures called “fruiting bodies.” Up to now, photosynthetic bacteria have not been reported among the δ-class. Menaquinones are the major electron carriers in the respiratory chain (Table 3). The major subgroups of the “Deltaproteobacteria” are: 1) the myxobacteria displaying gliding motility and forming multicellular fruiting bodies, which are often large enough to be observed with the aid of a hand lens or even by the naked eye (e.g., Chondromyces); 2) the bdellovibrios living as predators on other Gram-negative bacteria; 3) the dissimilatory sulfate- and sulfur-reducing bacteria (their genus names are mostly prefixed by Desulfo-); and 4) some synthrophic bacteria which ferment propionate (e.g., Syntropho-

22

K. Kersters et al.

bacter) or benzoate (e.g., Syntrophus) to acetate, CO2 and hydrogen syntrophically in coculture with hydrogen-consuming methanogens. Bdellovibrios and myxobacteria are strict aerobes, whereas the sulfate- and sulfur-reducers and the syntrophic bacteria are strictly anaerobic. One may speculate whether the bdellovibrios and the myxobacteria represent aerobic descendants of the sulfate- and sulfur-reducing bacteria. We consider here briefly the major phylogenetic groups of the “Deltaproteobacteria,” containing at present some 60 different genera and 160 species. Table 11 summarizes the names of taxa above the rank of genus within the “Deltaproteobacteria” according to the new edition of Bergey’s Manual of Systematic Bacteriology (Garrity, 2001). Myxobacteria The myxobacteria constitute a fascinating group of strictly aerobic, chemoorganotrophic, gliding, generally pigmented rodshaped cells, which under starvation conditions aggregate, forming multicellular and macroscopic fruiting bodies (see Dworkin and Kaiser [1993] and Dawid [2000]). Some of the cells within these fruiting bodies are often converted to resting stages called myxospores. The peculiar gliding movement of the vegetative cells is not restricted to myxobacteria as it occurs also amongst members of the genera Cytophaga and Flexibacter, both belonging to the CytophagaBacteroides-Flavobacterium phylum (now called “Bacteroidetes”; Garrity, 2001). New molecular and genetic insights concerning gliding motility have been gathered from studies on Myxococcus xanthus (Spormann, 1999). The myxobacteria are grouped in the order Myxococcales, which is composed of the following four families: Myxo-

coccaceae, Archangiaceae, Cystobacteraceae and the Polyangiaceae. Those fruiting myxobacteria whose rDNA sequence could be examined form a phylogenetically coherent group (Spröer et al., 1999; Fig. 1), consisting of two major phylogenetic lineages. One fairly homogeneous group is composed of the genera Myxococcus, Angiococcus, Archangium, Cystobacter, Melittangium and Stigmatella, whereas the second lineage is much more heterogeneous and is composed of three genera belonging to the family Polyangiaceae. Within the latter group, the genus Nannocystis seems to occupy the most separate position. The classification of the myxobacteria relies heavily on morphological features whose phylogenetic significance has been confirmed (Spröer et al., 1999). The myxobacteria occur in various habitats, such as soils and particularly on decaying organic material, including dung of herbivorous animals, rotting wood, and bark of living and dead trees. They obtain their nutrients primarily by causing lysis of other bacteria. The myxobacteria are mostly mesophilic, but some psychrophilic, acidophilic and alkaliphilic species have been described (Dawid, 2000). The sporangia of Chondromyces crocatus have been found to harbor a sphingobacteriumlike organism which was described as “Candidatus comitans” by Jacobi et al. (1997). Several myxobacteria produce potentially useful secondary compounds, such as antibiotics and cytostatic peptides (Reichenbach and Höfle, 1993; Reichenbach, 2001). The fruiting myxobacteria possess the most complex behavioral patterns and life cycles of all prokaryotes known so far and are therefore considered as a very interesting study object. They typically display social behavior expressed by collective food

Table 11. The “Deltaproteobacteria”: orders, families and number of genera.a Order “Desulfurellales” “Desulfovibrionales”

“Desulfobacterales”

“Desulfuromonadales”

“Synthrophobacterales” “Bdellovibrionales” Myxococcales

Family

Number of genera

“Desulfurellaceae” “Desulfovibrionaceae” “Desulfomicrobiaceae” “Desulfohalobiaceae” “Desulfobacteraceae” “Desulfobulbaceae” “Desulfoarculaceae” “Desulfuromonadaceae” “Geobacteraceae” “Pelobacteraceae” “Synthrophobacteraceae” “Synthrophaceae” “Bdellovibrionaceae” Myxococcaceae Archangiaceae Cystobacteraceae Polyangiaceae

2 3 1 3 12 4 4 2 1 2 4 2 3 2 1 3 3

a According to the second edition of Bergey’s Manual of Systematic Bacteriology (Garrity and Holt, 2001). Quotation marks are used for names which have not yet been validated (as of mid 2002).

Introduction to the Proteobacteria

uptake, cooperative motility (swarming), and social development (Crespi, 2001). Intercellular communication plays an important role in these phenomena. The analysis of the genome sequence of Myxococcus xanthus (9.5 Mb; twice as large as the E. coli genome) should enhance the understanding of the molecular and evolutionary basis of the peculiar life style of these myxobacteria. Bdellovibrio Group The bdellovibrios are small vibrioid prokaryotes living as predators on other Gram-negative bacteria. They are widespread in nature and have been isolated from soil, sewage, freshwater and marine environments. Wildtype bdellovibrios show a biphasic life cycle, alternating between an extracellular “attack-phase” flagellated form and an intracellular nonflagellated reproductive phase. After a bdellovibrio cell penetrates the wall of a Gramnegative prey cell (such as Pseudomonas or Erwinia), it performs a number of modifications on the outer membrane and the peptidoglycan of the invaded bacterium. The resulting spherical structure, consisting of the killed invaded cell and the developing bdellovibrio, is called a “bdelloplast.” The bdellovibrio cell grows and multiplies in the periplasmic space of the prey cell and digests the periplasmic and cytoplasmic contents of the invaded cell (for more details, see Jurkevitch, 2001). The parasitic life style of Bdellovibrio is reflected in the size of its genome, which is only half of that of E. coli. Genomic heterogeneity has been observed among the bdellovibrios (Seidler et al., 1972; Baer et al., 2000): the genera Bdellovibrio and Bacteriovorax constitute separate lineages among the “Deltaproteobacteria.” The 16S rDNA sequences of two other genera (Micavibrio and Vampirovibrio) have not been analyzed yet. The latter genus attacks cells of the green alga Chlorella. Sulfate- and Sulfur-reducing Bacteria Sulfate- and sulfur-reducing bacteria occur in various major phylogenetic lineages, such as the Archaea (e.g., Archaeglobus), the phylum of the thermophilic Thermodesulfobacteria, the order Clostridiales (the spore-forming Desulfotomaculum and Desulfosporosinus), and the “Deltaproteobacteria” (such as the genera Desulfovibrio, Desulfobacter and Desulfuromonas). The proteobacterial sulfate- and sulfur-reducing bacteria are mostly mesophilic and constitute a physiological group of an otherwise phylogenetically heterogeneous assemblage of bacteria (Fig. 1). The number of genera and species has significantly increased during the last decade: in 2002, the group encompassed some 40 validly published genera and some 120 species, as well as a number of not yet culturable bacteria which

23

emerge from molecular environmental studies (Akkermans et al., 1994). The proteobacterial sulfate- and sulfur-reducers are widespread in aquatic and humid terrestrial environments that become anoxic as a result of microbial decomposition. They are for the most part strictly anaerobic, performing the complete or partial oxidation of small organic molecules (e.g., lactate, pyruvate, acetate, and in a few cases, also alkanes and aromatic compounds) with oxidized S-compounds (such as sulfate or elemental sulfur) as terminal electron acceptors generating hydrogen sulfide. They play a crucial role in the sulfur cycle. Some sulfate reducers (e.g., Desulfonema and Desulfosarcina) can also grow autotrophically with hydrogen gas as electron donor and CO2 as sole carbon source. Multiheme cytochromes (of the c3 type) play a key role in electron-transport reactions in the periplasm of sulfate- and sulfur-reducing bacteria. Strictly sulfur-reducing bacteria occur among the genera Desulfuromonas, Desulfuromusa, Desulfurella and Hippea (Liesack and Finster, 1994; Miroshnichenko et al., 1999). Phylogenetically the sulfate- and sulfurreducing proteobacterial taxa display a significant diversity; at present at least five major phylogenetic lineages can be discerned within the “Deltaproteobacteria” (Fig. 1). To make the picture perhaps even more complex, not all the taxa belonging to these phylogenetic clusters metabolize with sulfate or sulfur as terminal electron acceptor. These five major rRNA clusters are the Desulfovibrio, Desulfuromonas, Desulfobacter, Synthrophobacter and Desulfurella groups. The Desulfovibrio rRNA group (encompassing the genera Desulfovibrio, Bilophila and Lawsonia) is phylogenetically related to the Desulfohalobium group, Desulfomicrobium and Desulfonatronum. Desulfovibrios occur in aquatic biotopes but are also common inhabitants of the intestines of mammals, including humans. Interestingly, Lawsonia intracellularis is an obligately intracellular bacterium occurring in the intestinal epithelial cells of pigs with proliferative enteropathy, a major disease affecting the economics of pig industries worldwide. Lawsonia does not reduce sulfate, but is clearly related to the Desulfovibrio group (McOrist et al., 1995). Similarly Bilophila wadsworthia is a common inhabitant of the human colon and has been associated with human appendicitis and was also reported as a common infective agent in neonatal pigs (McOrist et al., 2001). One species of Desulfomicrobium (D. orale) has been described as being involved in peridontal disease. The Desulfuromonas group is more related to the Pelobacter group, which is mainly composed of fermenting bacteria, such as Malonomonas

24

K. Kersters et al.

rubra, growing by the decarboxylation of malonate to acetate. Some of the sulfate-reducing bacteria can also utilize Fe3+ as electron acceptor, which is typical for the various Geobacter species, also belonging to this rRNA cluster. The Desulfobacter group is quite diverse as it contains 14 genera and is phylogenetically related to the Desulfobulbus and the Desulfomonile (or “Desulfoarculus”) groups. The Desulfobacter group harbors also some psychrophilic sulfatereducing bacteria such as Desulfofrigus and Desulfofaba, which were isolated from cold Arctic marine sediments (Knoblauch et al., 1999). Sulfate reducers occur also among the Syntrophobacter group (Fig. 1), which is composed of the syntrophs classified in the genus Syntrophobacter and the sulfate-reducers belonging to Desulforhabdus and Desulfovirga (Tanaka et al., 2000) together with the thermophilic marine genera Desulfacinum and Thermodesulforhabdus. The latter two genera form part of a subsurface microbial community; the former was isolated from hydrothermal vent systems and the latter from hot North Sea oil field water. Other syntrophs of the “Deltaproteobacteria” such as the benzoate-degrading Syntrophus species and the propionate-degrading Smithella belong to the Syntrophus group (Fig. 1). The genera Desulfurella and Hippea seem to occupy a separate position within the Proteobacteria; they are anaerobic, moderately thermophilic, sulfur-reducing bacteria, isolated from respectively terrestrial hot springs or submarine hot vents. Pending further studies, it is possible that in the future the Desulfurella group might be considered as a sixth (zeta) class among the Proteobacteria (Rainey et al., 1993).

The “Epsilonproteobacteria” This is the smallest and a more recently recognized line of descent within the Proteobacteria (6 genera and some 50 species). Compared to the multiple changes in the classification of the four other proteobacterial classes, the classification of the ε-class remained quite stable, because it is a fairly new group whose taxonomy was right from its recognition based on comparisons of 16S rDNAs. Up to now, photosynthetic bacteria have not been reported and key representatives are the genera Campylobacter and Helicobacter, encompassing enteropathogens for man and/or animals. Most species of the “Epsilonproteobacteria” are microaerophilic, chemoorganotrophic nonsaccharolytic spiral-shaped or curved bacteria, typically motile with a corkscrew-like motion by means of polar flagella. They obtain their energy mainly from amino acids or tricarboxylic acid cycle intermediates. Menaquinones are the characteristic respiratory quinones (Table 3). In

the 1980s, interest in campylobacters and related bacteria increased owing to the availability of adequate isolation and cultivation procedures. Some species require indeed fumarate, or formate plus fumarate, or hydrogen plus fumarate for growth in microaerobic conditions. Numerous new taxa were described, and the gastritiscausing Campylobacter pylori was transferred together with C. mustelae (isolated from the gastric mucosa of ferret) to the new genus Helicobacter by Goodwin et al. (1989). Some not yet culturable “Epsilonproteobacteria” have been reported as symbionts of shrimps and polychaetes. The complete genome sequences of Campylobacter jejuni (Parkhill et al., 2000) and Helicobacter pylori (Alm et al., 1999) have been determined. For recent reviews emphasizing taxonomic aspects of the major genera of the “Epsilonproteobacteria,” see Vandamme (2000), On (2001), and Solnick and Vandamme (2001), and also Wassenaar and Newell (2001). Table 12 summarizes the names of taxa above the rank of genus within the “Epsilonproteobacteria” according to the new edition of Bergey’s Manual of Systematic Bacteriology (Garrity, 2001). The Campylobacteraceae Campylobacter coli and C. jejuni are the most frequently identified causative agents of acute infective diarrhea in humans. Campylobacters can enter the food chain via raw milk and raw meat (particularly poultry). Other Campylobacter and Arcobacter species are involved in abortion and infectious infertility in various animals (e.g., sheep and cows), whereas some Campylobacter species are also found in large numbers in the periodontal pockets of diseased gums of man. Although primarily known as human and animal associated bacteria which are of relevance in food microbiology, arcobacters were shown to be abundantly present in certain environmental niches including water reservoirs, sewage, oil field communities, and certain saline environments. Their role in the environment is not well documented, but some of these organisms were shown to be sulfide oxiders (with the production of sulfur) and it has been suggested that they play a role in the sulfur cycle by reoxidizing sulfide formed by Table 12. The “Epsilonproteobacteria”: orders, families and number of genera.a Order “Campylobacterales”

Family

Number of genera

Campylobacteraceae “Helicobacteraceae”

4 2

a According to the second edition of Bergey’s Manual of Systematic Bacteriology (Garrity and Holt, 2001). Quotation marks are used for names which have not yet been validated (as of mid 2002).

Introduction to the Proteobacteria

microbial sulfate or sulfur reduction (Teske et al., 1996; Voordouw et al., 1996; Snaidr et al., 1997; Wirsen et al., 2002). The type species A. nitrofigilis occurs in the roots and the rhizosphere of salt marsh plants, where these bacteria fix nitrogen. The genus Sulfurospirillum harbors microaerophilic sulfur-reducing curved bacteria occurring in freshwater or marine surface sediments. Two recently described Sulfurospirillum species (S. barnesii and S. arsenophilum) can even use arsenate or selenate as electron acceptor (Stolz et al., 1999). The strictly anaerobic “Dehalospirillum multivorans” was isolated from activated sludge and can dechlorinate tetrachloroethene, a volatile man-made pollutant (Scholz-Muramatsu et al., 1995). This organism seems to be phylogenetically related to Sulfurospirillum. The Helicobacter Group The helicobacters constitute together with Wolinella a distinct phylogenetic lineage within the “Epsilonproteobacteria” (Fig. 1). Helicobacter pylori has received considerable attention for its role in gastric and duodenal ulcers and in gastric cancer in humans. Helicobacter encompasses at present some 20 species and more will likely be discovered when the stomach of other animals will be screened for bacteria similar to H. pylori. The phylogenetic and taxonomic position of various so-called “gastrospirilla” occurring in the stomachs of humans and animals will only be resolved when significant advances in the culture methods for these difficult bacteria are made. The pig and cattle strains now constitute two provisional candidate species: respectively Candidatus Helicobacter suis and Candidatus Helicobacter bovis (De Groote et al., 1999a; De Groote et al., 1999b; see also Table 13). Similarly, polyphasic investigations are needed in the group of “Flexispira rappini,” which is a provisional name for the spindle-shaped bacteria with periplasmic fibers and bipolar tufts of sheathed flagella, isolated from a variety of clinical and veterinary sources. The name “Flexispira rappini” refers to a characteristic morphotype shared by several distinct taxa and not to a distinct well-defined species (Dewhirst et al., 2001; Vandamme and On, 2001). Wolinella succinogenes (formerly Vibrio succinogenes), a commensal of the bovine rumen, belongs also to the Helicobacter group. This organism can grow on hydrogen as electron donor and fumarate as electron acceptor. Likely the full extent of the ecological diversity of the “Epsilonproteobacteria” still remains to be discovered: 16S rDNA gene sequence data of symbionts of shrimps (Rimicaris exoculata) and polychaete annelids (Alvinella pompejana) suggest that some “Epsilonproteobacteria” may occupy important niches in the habitats of deep-

25

sea hydrothermal vents, where they contribute to overall carbon and sulfur cycling at moderate thermophilic temperatures (Campbell et al., 2001; Corre et al., 2001). Recently, such a thermophilic sulfur-reducing bacterium isolated from the deep-sea hydrothermal vent polychaete, Alvinella pompejana, was allocated to a new genus, Nautilia (Miroshnichenko et al., 2002). Another remarkable member of the “Epsilonproteobacteria” is Thiovulum, whose phylogenetic relationship was shown by partial rDNA analysis (Romaniuk et al., 1987). The sulfuroxidizing Thiovulum possesses large ovoid cells with a diameter often reaching 25 µm. The cytoplasm contains orthorhombic sulfur inclusions, and the bacterium is considered as a chemolithotroph occurring in marine and freshwater environments, where sulfide-containing water and mud layers are in contact with overlaying oxygen-containing water. The environmental Thiomicrospira denitrificans belongs also to the “Epsilonproteobacteria,” whereas the other six Thiomicrospira species, including the type species T. pelophila, are affiliated to the Piscirickettsia group of the “Gammaproteobacteria” (Brinkhoff et al., 1999).

Symbiotic, Parasitic and Not-Yet Cultured Proteobacteria, and the Alphaproteobacterial Origin of Mitochondria The application of molecular biological methods to study the diversity of genes from environmental DNA without the need to isolate prokaryotic strains prior to molecular analyses has broadened immensely our insight into community structure and must be considered a quantum leap in microbial ecology (Hugenholz et al., 1998). Though problems are still associated with its ability to qualify and quantify individual partners of the community, the molecular approach has circumvented problems recognized with pure culture studies—the “great plate count anomaly,” as phrased by Staley and Konopka (1985). Hundreds of habitats and selected environmental sites have been subjected to the direct analysis of community structure by ribosomal rDNA sequencing and, more recently, of community function by the analysis of genes involved in metabolism (sulfur, nitrogen, methanogenesis, etc.) as well as degradation, bioremediation or heavy metal resistance. Symbiosis between prokaryotic and eukaryotic organisms seems to play a major role in evolution (Gray and Doolittle, 1982; Margulis, 1993; Margulis, 1996). A classical example is the symbiotic association between some diazotrophic

26

K. Kersters et al.

Table 13. Published Candidatus taxa among the Proteobacteria (as of mid 2002). Name of Candidatus Reference “Alphaproteobacteria” “Candidatus Liberibacter africanus” “Candidatus Liberibacter africanus subsp. capensis” “Candidatus Liberibacter asiaticus” “Candidatus Odyssella thessalonicensis” “Candidatus Xenohaliotis californiensis” “Betaproteobacteria” “Candidatus Procabacter acanthamoebae” “Gammaproteobacteria” “Candidatus Arsenophonus triatominarum” “Candidatus Blochmannia floridanus” “Candidatus Blochmannia herculeanus” “Candidatus Blochmannia rufipes” “Candidatus Phlomobacter fragariae” “Epsilonproteobacteria” “Candidatus Helicobacter bovis” “Candidatus Helicobacter suis”

Comments/host

References

Phloem-restricted bacterium associated with greening disease in citrus Phloem-restricted bacterium associated with leaf mottle symptoms in Calodendrum capense (Cape chestnut) Phloem-restricted bacterium associated with greening disease in citrus Endosymbiont of Acanthamoeba spp. Phylogenetically related to Caedibacter and Holospora Endosymbiont causing withering syndrome of Haliotis spp. (abalone). Belongs to the Rickettsiaceae

Jagoueix et al., 1994 Murray and Stackebrandt, 1995 Garnier et al., 2000

Endosymbiont of Acanthamoeba spp.

Horn et al., 2002

Endosymbiont of Triatoma infestans. Closely related to Arsenophonus nasoniae (the triatomine bug) Endosymbiont of Camponotus floridanus (carpenter ants) Endosymbiont of Camponotus herculeanus (carpenter ants) Endosymbiont of Camponotus rufipes (carpenter ants) Phloem-restricted bacterium associated with marginal chlorosis of strawberry

Hypsa and Dale, 1997

Occurs in pyloric part of the abomasum of cattle Occurs in stomach of pigs

“Alphaproteobacteria” (e.g., Rhizobium, Bradyrhizobium and Azorhizobium) and leguminous plants, where the bacteria reduce atmospheric nitrogen in root or stem nodules. Some Proteobacteria live as obligate intracellular symbionts or parasites in animals. To study the long-term history of symbiotic associations, the phylogenetic trees of hosts and their symbionts have been compared. Few data only are available, the most convincing study being the one on the endosymbionts of aphids. The topology of the symbiont (Buchnera aphidicola) tree is completely concordant with host phylogeny, based upon morphology (Moran et al., 1993). The fossil and biogeographic time points for the host phylogeny have been used to calibrate the 16S rDNA of the closely related endosymbionts (>8% similarity). The value of 1% fixed substitutions per 25 to 50 milllion years (Moran et al., 1993) is similar to the value of 1% per 50 million years determined by Ochman and Wilson (1987) on the basis of a broader range of nonobligate symbiotic relationships (e.g., Rhizobium/legumes; Photobacterium/fish; and enterobacteria/mammals) and 1% per 60 million years suggested by Stackebrandt (1995) for the past 500 years.

Jagoueix et al., 1994 Murray and Stackebrandt, 1995 Birtles et al., 2000

Friedman et al., 2000

Sauer et al., 2000 Sauer et al., 2000 Sauer et al., 2000 Zreik et al., 1998

De Groote et al., 1999a De Groote et al., 1999b

The advantage to the host of the association has been unraveled in only a few cases (e.g., the removal of hydrogen produced from hydrogenosomes of ciliates, provision of nutritional carbon to the host bivalves by sulfur-oxidizing gill symbionts [Dando and Southward, 1986] or of essential amino acids to aphids by their endosymbionts [Baumann et al., 1995; Baumann et al., 1998]). Molecular tools have also been used to elucidate the transmission route of symbionts in tropical lunicid bivalves (Gros et al., 1998) and deep-sea bivalves (Krueger et al., 1996) by application of the polymerase chain reaction (PCR) techniques in ovaries, testis and gill tissue. We can estimate that a considerable number of new proteobacterial taxa has not yet been detected or described, because suitable in vitro cultivation techniques do not yet exist for such endosymbiotic bacteria. However, DNA-based techniques allow the detection and visualization of such endosymbionts, and PCR amplification of their 16S rDNA by oligonucleotide probes (see Table 4), applied singly or in a combination of several probes, each representing a different taxonomic level, enables the determination of their phylogenetic position

Introduction to the Proteobacteria

(Manz et al., 1992; Amann et al., 1995; Amann and Ludwig, 2000). An intracellular lifestyle has obvious advantages for a suitably adapted prokaryote, and the spread of intracellular bacteria seems to be guaranteed in hosts such as blood sucking insects. Hence, it is not surprising that intracellular bacteria have evolved in different phylogenetic groups. Classical examples among the Proteobacteria are Coxiella, Rickettsiella and the endosymbionts of aphids belonging to the genus Buchnera (all “Gammaproteobacteria”) and various members of the Rickettsiaceae and allied groups (e.g., Holospora and Caedibacter), belonging to the “Alphaproteobacteria.” Related to the Rickettsiaceae are the wolbachiae, which are symbionts of invertebrate hosts, affecting the reproduction of the host by induction of thelytokous parthenogenesis (i.e., male killing or feminization). The wolbachiae infect the reproductive cells of various insects and have thus the potential to be vertically transmitted by cytoplasmic inheritance. More details concerning intracellular prokaryotes can be found in the introductory chapter by Fredricks (Introduction to the Rickettsiales and other Intracellular Prokaryotes in this Volume) and some examples of symbiotic and not yet cultured Proteobacteria are shown in Table 14, together with their phylogenetic affiliation.

27

Most of these endosymbiotic bacteria cannot yet be cultivated and illustrate that only a very limited proportion of the bacterial species present in various habitats is actually known. Although a great part of the rDNA sequences of uncultured bacteria is available in public databases, no formal phenotypic description nor species names can be given for such organisms, and type strains cannot be made publicly available. Some of these uncultured bacteria have been partially described and provisionally allocated to the category of Candidatus (Murray and Schleifer, 1994; Murray and Stackebrandt, 1995). The proteobacterial Candidatus taxa together with some additional information are listed in Table 13 (see also the List of Bacterial Names with Standing in Nomenclature website). The mitochondrion of eukaryotic cells can be considered as the ultimate prokaryotic endosymbiont. Like chloroplasts, mitochondria possess unique genomes that are distinct from the nuclear genomes of their associated eukaryotic cells, and mitochondrial ribosomes are clearly of the prokaryotic type and related to the “Alphaproteobacteria” (Lang et al., 1999). Among the many bacterial genomes that have now been sequenced, that of Rickettsia prowazekii (1.1 Mb) is most closely related to the mitochondrial genome (Andersson et al., 1998). Rickettsia prowazekii is the agent of epidemic

Table 14. Examples of symbiotic Proteobacteria and their phylogenetic affiliation. Host species Muscidifurax uiniraptor Bangasternus orientalis Rhinocyllus conicus Trichogramma deion Nasonia vitripennis Crassostrea virginica Dysmicoccus neobrevipes Acromyrmex octospinosus Antonina crawii Solemya velum Heliothis virescens Bathymodiolus thermophilus Calyptogena elongata Calyptogena magnifica Vesicomya cordata Anodontia phillipiana Codakia orbicularis Thyasira flexuosa Lucina floridana Riftia pachyptila Anomalops katoptron Cryptosaras couesi Sitophilus zeamais Acyrthosiphon sp. Camponotus floridanus Glossina pallidipes

Host trivial name

Phylogenetic affiliation of symbiont

Reference/EMBL accession number

House fly Coleoptera, weevil Coleoptera, weevil Parasite wasp of Lepidoptera Parasitic wasp Eastern Oyster Homoptera, mealy bug Leave cutting ant Bamboo pseudococcoid, Homoptera Bivalve Moth, tobacco budworm Bivalve Bivalve Bivalve Bivalve Bivalve Bivalve Bivalve Bivalve Tube worm Deep sea anglerfish Deep sea anglerfish Coleoptera, maize weevil Pea aphid Giant ants Tsetse fly

α-class α-class α-class α-class α-class α-class β-class β-class β, γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class γ-class

Stouthamer et al., 1993 M85266 M85267 Stouthamer et al., 1993 Gherna et al., 1991 Boettcher et al., 2000 Munson et al., 1992 AF459796 Fukatsu et al., 2000 M90415 L22481 Distel et al., 1988 L25719 Distel et al., 1988 L25713 L25711 Distel et al., 1988 Distel and Wood, 1992 L25707 Distel et al., 1988 Z19081 Haygood et al., 1992 M85270 Unterman et al., 1989 Schröder et al., 1996 Beard et al., 1993

28

K. Kersters et al.

louse-borne typhus in humans and multiplies in eukaryotic cells only. Most likely the genome types of mitochondria and rickettsiae have shared a common free-living alphaproteobacterial ancestor, which through genome reduction evolved via two descendent lineages (Gray, 1998; Gray et al., 1999). Studies on mitochondria of yeast reveal that the number of alphaproteobacterial descendents in the mitochondrial proteome is surprisingly small, and that a large number of novel mitochondrial genes likely originated from the eukaryotic nuclear genome complementing the remaining genes from the bacterial ancestor (Karlberg et al., 2000; Kurland and Andersson, 2000). Further genomic and proteomic analyses of other members of the “Alphaproteobacteria,” such as Bradyrhizobium and Rhodobacter, may yield more information concerning the metabolic versatility of the protomitochondrion. Whether the origin of hydrogenosomes—the ATP-producing organelles of many anaerobic protists—is related to that of mitochondria remains to be investigated in more detail (Andersson and Kurland, 1999; Dyall et al., 2000).

Genome Analysis The G+C content of genomic DNA of the Proteobacteria varies from 30 mol% (e.g., in Campylobacter and Rickettsia) to 70% (e.g., in Alcaligenes), indicating that almost the entire span of mol% G+C variation among living microorganisms is covered. Even within each of the proteobacterial subclasses, variation in G+C content of the genome is very large. Similarly the genome sizes of various Proteobacteria differ considerably. Commensals and free-living prokaryotic strains have a larger genome, ranging from 9.4 Mb (Myxococcus xanthus) to about 2.5 Mb (Pasteurella multocida), than that of obligate symbiotic and parasitic strains, e.g., Buchnera aphidicola (0.64 Mb) and Rickettsia prowazekii (1.1 Mb). Unexpectedly, the genome size of phylogenetically closely related species and even strains of the same species can differ significantly, as shown for three species of Yersinia (3.8–4.9 Mb) and various strains of E. coli (4.6–5.5 Mb) and Burkholderia cepacia (4.7–8.1 Mb; Lessie et al., 1996). Most Proteobacteria studied contain a single circular chromosome, but the presence of multiple chromosomes has been reported, particularly among the “Alphaproteobacteria”: two different circular chromosomes in Rhodobacter sphaeroides (Suwanto and Kaplan, 1989) and Brucella melitensis (Michaux-Charachon et al., 1997); three circular chromosomes in Rhizobium

meliloti (Sobral et al., 1991) and Burkholderia cepacia (Cheng and Lessie, 1994). Another unconventional organization has been detected in several strains of Agrobacterium tumefaciens, where the larger chromosome is circular (3.0 Mb), whereas the smaller (2.1 Mb) is linear (Jumas-Bilak et al., 1998). It should be noted that not all strains of Brucella melitensis or all species of the genus Rhodobacter contain multiple chromosomes, and the evolutionary significance of these peculiar genomic organizations remains to be unraveled (Jumas-Bilak et al., 1998). A multitude of genes of various Proteobacteria have been sequenced; genetic and genomic aspects of E. coli, various Pseudomonas species, Rhizobium and many animal and human pathogenic Proteobacteria have been or are being studied in detail, and more importantly in recent years, considerable progress has been made in the sequencing of whole genomes of an increasing number of Proteobacteria. The first wholegenome sequence of a free-living microorganism was published in 1995 by Fleishmann et al. for the gammaproteobacterium Haemophilus influenzae (1.83 Mb). At present (mid 2002), the sequences have been completed for at least 9 α-, 3 β- (Bordetella and Neisseria), 17 γ- and 3 ε(Campylobacter and Helicobacter) class members (sequenced genera are indicated by footnote “a” in Table 3; see also The Institute for Genomic Research website). Sequencing of a great number of proteobacterial genomes is in progress (see footnote “b” in Table 3 and The Institute for Genomic Research website, National Center for Biotechnological Information website, ([DOE Joint Genome Institute website] http://www.jgi. doe.gov/JGI_microbial/html and [GOLD Genome OnLine Database website] wit.integratedgenomics.com/GOLD/) and most of it pertains to bacteria of clinical or biotechnological importance. The sequencing projects will elucidate regulatory and structural functions of newly discovered genes and will also yield significant insights into the mechanisms of pathogenicity, bacterial photosynthesis, phylogeny and evolution.

Final Considerations With the publication of the Approved Lists of Bacterial Names in 1980, the number of genera and species presently belonging in the Proteobacteria was approximately 130 and 500, respectively. Until 2002, the number of validly described genera and species increased to respectively about 460 and 1600, making the phylum Proteobacteria a heavily populated section

Introduction to the Proteobacteria

of the phylogenetic bacterial tree. Indeed, in terms of number of genera, the Proteobacteria encompass more than 40% of all prokaryotic genera. Recent insights into the community structure of as-yet uncultivated prokaryotes reveal that the vast majority of taxa have not yet been described. This is true not only for open environments, such as the open ocean (Giovannoni et al., 1990), soil (Liesack and Stackebrandt, 1992; Felske et al., 1999), deep subsurface (Chandler et al., 1997), waste treatment plants and bioreactors (Bond et al., 1995; Juretschko et al., 2002), and sediments of oceans and lakes (Bowman et al., 2000; Urakawa et al., 2000; Brambilla et al., 2001), but also for the microflora of eukaryotic hosts. Particularly the not-yet culturable Proteobacteria, the endosymbionts and those which are difficult to grow in axenic culture such as the helicobacters and allied taxa, are a goldmine for further studies on bacterial diversity. The most promising tools to explain the role of Proteobacteria in global cycling of gases and chemicals are: 1) for elucidating the phylogenetic structure of the community, DNA fragment electrophoresis, sequence analysis of universal and taxon-specific genes, and their identification in situ by FISH hybridization, and 2) for determining the physiological capacities of its members, identification of housekeeping genes by FISH techniques and by microautoradiography (Ouverney and Fuhrman, 1999; Lee et al., 1999). Horizontal transfer of genes (Klein et al., 2001) will shed light on the possible evolutionary history of the various large groups within Proteobacteria by tracing the origin of genes to the ancestors of recent species that are phylogenetically related to other prokaryotic phyla. The number of full genome sequences will continue to rise, providing microbiologists with an unparalleled wealth of information for scientific exploitation to the benefit of clinical, environmental, evolutionary and general microbiology. It is likely that the increasing number of genera and species as well as comparative studies on other semantic macromolecules will challenge the present five major subdivisions of the Proteobacteria: as the phylogenetic radiation of the proteobacterial lineages will become more complex, boundaries between the five major subgroups will become vague and the subgroups more ambiguous. On the other hand, information on orthologous genes other than ribosomal rRNA genes will provide systematicists with sets of molecules to be included in future studies on multi-locus sequence analysis. As recommended, this information will be used in the description of the taxon species (Stackebrandt et al., 2002) and possibly of higher taxa. Molecular studies and culture attempts will continue to go hand in hand. The scientific chal-

29

lenge in the near future will also include molecular determination of metagenomes from proteobacteria of selected sites as well as the cultivation of hitherto uncultured proteobacterial symbiotic and pathogenic organisms.

Literature Cited Achenbach, L. A., U. Michaelidou, R. A. Bruce, J. Frymann, and J. D. Coates. 2001. Dechloromonas agitata gen. nov., sp. nov. and Dechlorosoma suillum gen. nov. sp. nov., two novel environmentally dominant (per)chloratereducing bacteria and their phylogenetic position. Int. J. Syst. Evol. Microbiol. 51:527–533. Achtman, M. 1998. Microevolution during epidemic spread of Neisseria meningitidis. Electrophoresis 19:593–596. Akkermans, A. D. L., M. S. Mirza, H. J. M. Harmsen, H. J. Blok, P. R. Herron, A. Sessitsch, and W. M. Akkermans. 1994. Molecular ecology of microbes: A review of promises, pitfalls and true progress. FEMS Microbiol. Rev. 15:185–194. Alm, R. A., L. S. L. Ling, D. T. Moir et al. 1999. Genomicsequence comparison of two unrelated isolates of the human gastric pathogen Helicobacter pylori. Nature 397:176–180. Amann, R. I., W. Ludwig, and K.-H. Schleifer. 1995. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59:143–169. Amann, R., and W. Ludwig. 2000. Ribosomal RNA-targeted nucleic acid probes for studies in microbial ecology. FEMS Microbiol. Rev. 24:555–565. Ambler, R. P., J. Hermoso, M. Daniel et al. 1979. Cytochrome-c2 sequence variation among the recognized species of purple non-sulfur photosynthetic bacteria. Nature 278:659–660. Andersson, S. G. E., A. Zomorodipour, J. O. Andersson et al. 1998. The genome sequence of Rickettsia prowazekii and the origin of mitochondria. Nature 396:133–140. Andersson, S. G. E., and C. G. Kurland. 1999. Origins of mitochondria and hydrogenosomes. Curr. Opin. Microbiol. 2:535–541. Anzai, Y., H. Kim, J.-Y. Park, H. Wakabayashi, and H. Oyaizu. 2000. Phylogenetic affiliation of the pseudomonads based on 16S rRNA sequence. Int. J. Syst. Evol. Microbiol. 50:1563–1589. Aragno, M., and H. G. Schlegel. 1992. The mesophilic hydrogen-oxidizing (Knallgas) bacteria. In: A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York, NY. 344-384. Arahal, D. R, W. Ludwig, K.-H. Schleifer, and A. Ventosa. 2002. Phylogeny of the family Halomonadaceae based on 23S and 16S rDNA sequence analysis. Int. J. Syst. Evol. Microbiol. 52:241–249. Auling, G. 1992. Polyamines, biomarker for taxonomy and ecology of phytopathogenic and other bacteria belonging to the Proteobacteria. Belgian J. Bot. 125:203–209. Baer, M. L., J. Ravel, J. Chun, R. T. Hill, and H. N. Williams. 2000. A proposal for the reclassification of Bdellovibrio stolpii and Bdellovibrio starrii into a new genus Bacteriovorax gen. nov. as Bacteriovorax stolpii comb. nov. and Bacteriovorax starrii comb. nov., respectively. Int. J. Syst. Evol. Microbiol. 50:219–224.

30

K. Kersters et al.

Baumann, P., L. Baumann, C.-Y. Lai, D. Roubakhsh, N. A. Moran, and M. A. Clark. 1995. Genetics, physiology, and evolutionary relationships of the genus Buchnera: Intracellular symbionts of aphids. Ann. Rev. Microbiol. 49:55–94. Baumann, P., L. Baumann, M. A. Clark, and M. L. Thao. 1998. Buchnera aphidicola: The endosymbiont of aphids. ASM News 64:203–209. Beard, C. B., S. L. O’Neill, P. Mason, L. Mandelco, C. R. Woese, R. B. Tesh, F. F. Richards, and S. Aksoy. 1993. Genetic transformation and phylogeny of bacterial symbionts from tsetse. Insect Molec. Biol. 1:123–131. Birch, R. G. 1997. Plant transformation: problems and strategies for practical application. Ann. Rev. Plant Physiol. 48:297–326. Birtles, R. J., T. J. Rowbotham, R. Michel, D. G. Pitcher, B. Lascola, S. Alexiou-Daniels, and D. Raoult. 2000. “Candidatus Odyssella thessalonicensis” gen. nov., sp nov., an obligate intracellular parasite of Acanthamoeba species. Int. J. Syst. Evol. Microbiol. 50:63–72. Boettcher, K. J., B. J. Barber, and J. T. Singer. 2000. Additional evidence that juvenile oyster disease is caused by a member of the Roseobacter group and colonization of nonaffected animals by Stappia stellulata-like strains. Appl. Environ. Microbiol. 66:3924–3930. Bond, P. L., P. Hugenholtz, J. Keller, and L. L. Blackall. 1995. Bacterial community structures of phosphate-removing and non-phosphate-removing activated sludges from sequencing batch reactors. Appl. Environ. Microbiol. 61:1910–1916. Bowman, J. P., L. I. Sly, and E. Stackebrandt. 1995. The phylogenetic position of the family Methylococcaceae. Int. J. Syst. Bacteriol. 45:182–185. Bowman, J. P., J. Cavanagh, J. J. Austin, and K. Sanderson. 1996. Novel Psychrobacter species from Antarctic ornithogenic soils. Int. J. Syst. Bacteriol. 46:841–848. Bowman, J. P., S. M. Rea, S. A. McCammon, and T. A. McMeekin. 2000. Diversity and community structure within anoxic sediment from marine salinity meromictic lakes and a coastal meromictic marine basin, Vestfold Hills, Eastern Antarctica. Environ. Microbiol. 2:227–237. Brambilla, E., H. Hippe, A. Hagelstein, B. J. Tindall, and E. Stackebrandt. 2001. 16S rDNA diversity of cultured and uncultured prokaryotes of a mat sample from Lake Fryxell, McMurdo Dry Valleys, Antartica. Extremophiles 5:23–33. Brenner, D. J., S. P. O’Connor, H. H. Winkler, and A. G. Steigerwalt. 1993. Proposals to unify the genera Bartonella and Rochalimaea, with descriptions of Bartonella quintana comb. nov., Bartonella vinsonii comb. nov., Bartonella henselae comb. nov., and Bartonella elizabethae comb. nov., and to remove the family Bartonellaceae from the order Rickettsiales. Int. J. Syst. Bacteriol. 43:777–786. Brinkhoff, T., G. Muyzer, C. O. Wirsen, and J. Kuever. 1999. Thiomicrospira kuenenii sp. nov. and Thiomicrospira frisia sp. nov., two mesophilic obligately chemolithoautotrophic sulfur-oxidizing bacteria isolated from an intertidal mud flat. Int. J. Syst. Bacteriol. 49:385–392. Broda, E. 1970. The evolution of bioenergetic processes. Prog. Biophys. Molec. Biol. 21:145–208. Broda, E. 1975. The Evolution of Bioenergetic Processes. Pergamon Press. Oxford, UK. Buchanan, R. E., and N. E. Gibbons (Eds.). 1974. Bergey’s Manual of Determinative Bacteriology, 8th ed. Williams and Wilkins. Baltimore, MD. 1–1246.

Burkholder, W. H. 1950. Sour skin, a bacterial rot of onion bulbs. Phytopathol. 40:115–117. Busse, J., and G. Auling. 1988. Polyamine pattern as a chemotaxonomic marker withing the Proteobacteria. Syst. Appl. Microbiol. 11:1–8. Busse, H. J., E. B. M. Denner, and W. Lubitz. 1996. Classification and identification of bacteria: Current approaches to an old problem—overview of methods used in bacterial systematics. J. Biotechnol. 47:3–38. Byng, G. S., J. L. Johnson, R. J. Whitaker, R. L. Gherna, and R. A. Jensen. 1983. The evolutionary pattern of aromatic amino acid biosynthesis and the emerging phylogeny of pseudomonad bacteria. J. Molec. Evol. 19:272–282. Campbell, B. J., C. Jeanthon, J. E. Kostka, G. W. Luther, and S. C. Cary. 2001. Growth and phylogenetic properties of novel bacteria belonging to the epsilon subdivision of the Proteobacteria enriched from Alvinella pompejana and deep-sea hydrothermal vents. Appl. Environ. Microbiol. 67:4566–4572. Cavalier-Smith, T. 2002. The neomuran origin of archaebacteria, the negibacterial root of the universal tree and bacterial megaclassification. Int. J. Syst. Evol. Microbiol. 52:7–76. Chandler, D. P., S. M. Li, C. M. Spadoni, G. R. Drake, D. L. Balkwill, J. K. Fredrickson, and F. J. Brockman. 1997. A molecular comparison of culturable aerobic heterotrophic bacteria and 16S rDNA clones derived from a deep subsurface sediment. FEMS Microbiol. Ecol. 23:131–144. Chen, W.-M., S. Laevens, T.-M. Lee, T. Coenye, P. De Vos, M. Mergeay, and P. Vandamme. 2001. Ralstonia taiwanensis sp. nov., isolated from root nodules of Mimosa species and sputum of a cystic fibrosis patient. Int. J. Syst. Evol. Microbiol. 51:1729–1735. Cheng, H.-P., and T. G. Lessie. 1994. Multiple replicons constituting the genome of Pseudomonas cepacia 17616. J. Bacteriol. 176:4034–4042. Coenye, T., P. Vandamme, J. R. W. Govan, and J. J. LiPuma. 2001. Taxonomy and identification of the Burkholderia cepacia complex. J. Clin. Microbiol. 39:3427–3436. Collins, M. D., and D. Jones. 1981. Distribution of isoprenoid quinone structural types in bacteria and their taxonomic implications. Microbiol. Rev. 45:316–354. Collins, M. D., and F. Widdel. 1986. Respiratory quinones of sulfate-reducing and sulfur-reducing bacteria: A systematic investigation. Syst. Appl. Microbiol. 8:8–18. Colwell, R. R. 1970. Polyphasic taxonomy of the genus Vibrio: Numerical taxonomy of Vibrio cholerae, Vibrio parahaemolyticus, and related Vibrio species. J. Bacteriol. 104:410–433. Corre, E., A.-L. Reysenbach, and D. Prieur. 2001. Epsilonproteobacterial diversity from a deep-sea hydrothermal vent on the Mid-Atlantic Ridge. FEMS Microbiol. Lett. 205:329–335. Crespi, B. J. 2001. The evolution of social behaviour in microorganisms. Trends Ecol. Evol. 16:178–183. Dando, P. R., and A. J. Southward. 1986. Chemoautotrophy in bivalve mollusks of the genus Thyasira. J. Mar. Biol. Assoc. UK 66:915–929. Dauga, C., M. Gillis, P. Vandamme, E. Ageron, F. Grimont, K. Kersters, C. De Mahenge, Y. Peloux, and P. A. D. Grimont. 1993. Balneatrix alpica gen. nov., sp. nov. a bacterium associated with pneumonia and meningitis in a spa therapy centre. Res. Microbiol. 144:35–46. Dawid, W. 2000. Biology and global distribution of myxobacteria in soils. FEMS Microbiol. Rev. 24:403–427.

Introduction to the Proteobacteria Dedysh, S. N., V. N. Khmelenina, N. E. Suzina, Y. A. Trotsenko, J. D. Semrau, W. Liesack, and J. M. Tiedje. 2002. Methylocapsa acidiphila gen. nov., sp. nov., a novel methane-oxidizing and dinitrogen-fixing acidophilic bacterium from sphagnum bog. Int. J. Syst. Evol. Microbiol. 52:251–261. De Groote, D., L. J. Van Doorn, R. Ducatelle, A. Verschuuren, F. Haesebrouck, W. G. V. Quint, K. Jalava, and P. Vandamme. 1999a. “Candidatus Helicobacter suis”, a gastric helicobacter from pigs, and its phylogenetic relatedness to other gastrospirilla. Int. J. Syst. Bacteriol. 49:1769–1777. De Groote, D., L. J. Van Doorn, R. Ducatelle, A. Verschuuren, K. Tilmant, W. G. V. Quint, F. Haesebrouck, and P. Vandamme. 1999b. Phylogenetic characterization of “Candidatus Helicobacter bovis,” a new gastric helicobacter in cattle. Int. J. Syst. Bacteriol. 49:1707–1715. de Lajudie, P., A. Willems, G. Nick, F. Moreira, F. Molouba, B. Hoste, U. Torck, M. Neyra, M. D. Collins, K. Lindstrom, B. Dreyfus, and M. Gillis. 1998. Characterization of tropical tree rhizobia and description of Mesorhizobium plurifarium sp. nov. Int. J. Syst. Bacteriol. 48:369–382. De Ley, J., M. Gillis, and J. Swings. 1984. Family VI: Acetobacteraceae Gillis and De Ley 1980, 23. In: N. R. Krieg and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 1:267–277. De Ley, J., P. Segers, K. Kersters, W. Mannheim, and A. Lievens. 1986. Intrageneric and intergeneric similarities of the Bordetella ribosomal ribonucleic-acid cistrons: Proposal for a new family, Alcaligenaceae. Int. J. Syst. Bacteriol. 36:405–414. De Ley, J., W. Mannheim, R. Mutters, K. Piechulla, R. Tytgat, P. Segers, M. Bisgaard, W. Frederiksen, K.-H. Hinz, and M. Vanhoucke. 1990. Inter- and intrafamilial similarities of rRNA cistrons of the Pasteurellaceae. Int. J. Syst. Bacteriol. 40:126–137. De Ley, J. 1992. The Proteobacteria: Ribosomal RNA cistron similarities and bacterial taxonomy. In: A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York, NY. 2111–2140. De Vos, P., and J. De Ley. 1983. Intra- and intergeneric similarities of Pseudomonas and Xanthomonas ribosomal ribonucleic acid cistrons. Int. J. Syst. Bacteriol. 33:487– 509. De Vos, P., M. Goor, M. Gillis, and J. De Ley. 1985. Ribosomal ribonucleic acid cistron similarities of phytopathogenic Pseudomonas species. Int. J. Syst. Bacteriol. 35:169–184. De Vos, P., A. Van Landschoot, P. Segers, R. Tytgat, M. Gillis, M. Bauwens, R. Rossau, M. Goor, B. Pot, K. Kersters, P. Lizzaraga, and J. De Ley. 1989. Genotypic relationships and taxonomic localization of unclassified Pseudomonas and Pseudomonas-like strains by deoxyribonucleic acid:ribosomal ribonucleic acid hybridizations. Int. J. Syst. Bacteriol. 39:35–49. Dewhirst, F. E., B. J Paster, S. La Fontaine, and J. I. Rood. 1990. Transfer of Kingella indologenes (Snell and Lapage 1976) to the genus Suttonella gen. nov. as Suttonella indologenes comb. nov.; transfer of Bacteroides nodosus (Beveridge 1941) to the genus Dichelobacter gen. nov. as Dichelobacter nodosus comb. nov.; and assignment of the genera Cardiobacterium, Dichelobacter, and Suttonella to Cardiobacteriaceae fam. nov.

31

in the gamma division of Proteobacteria on the basis of 16S rRNA sequence comparisons. Int. J. Syst. Bacteriol. 40:426–433. Dewhirst, F. E., J. G. Fox, E. N. Mendes, B. J. Paster, C. E. Gates, C. A. Kirkbride, and K. A. Eaton. 2000. “Flexispira rappini” strains represent at least 10 Helicobacter taxa. Int. J. Syst. Evol. Microbiol. 50:1781–1787. Dickerson, R. E. 1980. Cytochrome-c and the evolution of energy metabolism. Scientific Am. 242(3):99–110. Distel, D. L., D. J. Lane, G. J. Olsen, S. J. Giovannoni, B. Pace, N. R. Pace, D. A. Stahl, and H. Felbeck. 1988. Sulfuroxidizing bacterial endosymbionts: Analysis of phylogeny and specificity by 16S rRNA sequences. J. Bacteriol. 170:2506–2510. Distel, D. L., and A. P. Wood. 1992. Characterization of the gill symbiont of Thyasira flexuosa (Thyasiridae: Bivalvia) by use of polymerase chain reaction and 16S rRNA sequence analysis. J. Bacteriol. 174:6317–6320. Drozanski, W. J. 1991. Sarcobium lyticum gen. nov., sp. nov., an obligate intracellular bacterial parasite of small freeliving amoebae. Int. J. Syst. Bacteriol. 41:82–87. Dumler, J. S., A. F. Barbet, C. P. J. Bekker, G. A. Dasch, G. H. Palmer, S. C. Ray, Y. Rikihisa, and F. R. Rurangirwa. 2001. Reorganization of genera in the families Rickettsiaceae and Anaplasmataceae in the order Rickettsiales: Unification of some species of Ehrlichia with Anaplasma, Cowdria with Ehrlichia and Ehrlichia with Neorickettsia, descriptions of six new species combinations and designation of Ehrlichia equi and “HGE agent” as subjective synonyms of Ehrlichia phagocytophila. Int. J. Syst. Evol. Microbiol. 51:2145–2165. Dunlap, P. V., and K. Kita-Tsukamoto. 2001. Luminous bacteria. In: M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer, and E. Stackebrandt (Eds.) The Prokaryotes, 3rd ed. Springer-Verlag. New York, NY. Dworkin, M., and D. Kaiser (Eds.). 1993. Myxobacteria II. American Society for Microbiology. Washington, DC. Dyall, S. D., C. M. Koehler, M. G. Delgadillo-Correa et al. 2000. Presence of a member of the mitochondrial carrier family in hydrogenosomes: Conservation of membranetargeting pathways between hydrogenosomes and mitochondria. Molec. Cell. Biol. 20:2488–2497. Ehrenberg, C. G. 1836. Vorläufige Mitteilungen über das wirkliche Vorkommen fossiler Infusorien und ihre große Verbreitung. Poggendorf’s Ann. Phys. Chem. 38:213– 227. Falah, M., and R. S. Gupta. 1994. Cloning of the HSP70 (DNAK) genes from Rhizobium meliloti and Pseudomonas cepacia: Phylogenetic analyses of mitochondrial origin based on a highly conserved protein sequence. J. Bacteriol. 176:7748–7753. Felske, A., A. Wolterink, R. Van Lis, W. M. De Vos, and A. D. L. Akkermans. 1999. Searching for predominant soil bacteria: 16S rDNA cloning versus strain cultivation. FEMS Microbiol. Ecol. 30:137–145. Finkmann, W., K. Altendorf, E. Stackebrandt, and A. Lipski. 2000. Characterization of N2O-producing Xanthomonas-like isolates from biofilters as Stenotrophomonas nitritireducens sp. nov., Luteimonas mephitis gen. nov., sp. nov. and Pseudoxanthomonas broegbernensis gen. nov., sp. nov. Int. J. Syst. Evol. Microbiol. 50:273–282. Fleischmann, R. D., M. D. Adams, O. White et al. 1995. Whole-genome random sequencing and assembly of Haemophilus influenzae Rd. Science 269:496–512. Fox, G. E., K. J. Pechman, and C. R. Woese. 1977. Comparative cataloguing of 16S ribosomal nucleic acid: Molec-

32

K. Kersters et al.

ular approach to prokaryotic systematics. Int. J. Syst. Bacteriol. 27:44–57. Fox, G. E., E. Stackebrandt, R. B. Hespell, J. Gibson, J. Maniloff, T. A. Dyer, R. S. Wolfe, W. E. Balch, R. S. Tanner, L. J. Magrum, L. B. Zablen, R. Blakemore, R. Gupta, L. Bonen, B. J. Lewis, D. A. Stahl, K. R. Luehrsen, K. N. Chen, and C. R. Woese. 1980. The phylogeny of prokaryotes. Science 209:457–463. Fredricks, D. N. 2001. Introduction to the Rickettsiales and other intracellular prokaryotes. In: M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer and E. Stackebrandt (Eds.) The Prokaryotes. Springer-Verlag. New York, NY. Friedman, C. S., K. B. Andree, K. A. Beauchamp, J. D. Moore, T. T. Robbins, J. D. Shields, and R. P. Hedrick. 2000. “Candidatus Xenohaliotis californiensis,” a newly described pathogen of abalone, Haliotis spp., along the west coast of North America. Int. J. Syst. Evol. Microbiol. 50:847–855. Fuentes-Ramirez, L. E., R. Bustillos-Cristales, A. TapiaHernandez, T. Jiménez-Salgado, E. T. Wang, E. Martinez-Moreno, and J. Caballero-Mellado. 2001. Novel nitrogen-fixing acetic acid bacteria, Gluconacetobacter johannae sp. nov. and Gluconacetobacter azotocaptans sp. nov., associated with coffee plants. Int. J. Syst. Evol. Microbiol. 51:1305–1314. Fukatsu, T., N. Nikoh, R. Kawai, and R. Koga. 2000. The secondary endosymbiotic bacterium of the pea aphid Acyrthosiphon pisum (Insecta: Homoptera). Appl. Environ. Microbiol. 66:2748–2758. Garnier, M., S. Jagoueix-Eveillard, P. R. Cronje, H. F. Le Roux, and J. M. Bové. 2000. Genomic characterization of a liberibacter present in an ornamental rutaceous tree, Calodendrum capense, in the Western Cape province of South Africa. Proposal of “Candidatus Liberibacter africanus subsp. capensis.” Int. J. Syst. Evol. Microbiol. 50:2119–2125. G. Garrity (Ed.). 2001. Bergey’s Manual of Systematic Bacteriology, 2nd ed. Springer-Verlag. New York, NY. 1:1– 721. Gaunt, M. W., S. L. Turner, L. Rigottier-Gois, S. A. Lloyd-Macgilp, and J. P. W. Young. 2001. Phylogenies of atpD and recA support the small subunit rRNA-based classification of rhizobia. Int. J. Syst. Evol. Microbiol. 51: 2037–2048. Gest, H., and J. L. Favinger. 1983. Heliobacterium chlorum, an anoxygenic brownish-green photosynthetic bacterium containing a new form of bacteriochlorophyll. Arch. Microbiol. 136:11–16. Gherna, R. L., J. H. Werren, W. Weisburg, R. Cote, C. R. Woese, L. Mandelco, and D. J. Brenner. 1991. Arsenophonus nasoniae gen. nov., sp. nov., the causative agent of the son-killer trait in the parasitic wasp Nasonia vitripennis. Int. J. Syst. Bacteriol. 41:563–565. Gibson, J., E. Stackebrandt, L. B. Zablen, R. Gupta, and C. R. Woese. 1979. Phylogenetic analysis of the purple photosynthetic bacteria. Curr. Microbiol. 3:59–64. Gillis, M., and J. De Ley. 1980. Intrageneric and intergeneric similarities of the ribosomal ribonucleic-acid cistrons of Acetobacter and Gluconobacter. Int. J. Syst. Bacteriol. 30:7–27. Giovannoni, S. J., T. B. Britschgi, C. L. Moyer, and K. G. Field. 1990. Genetic diversity in Sargasso Sea bacterioplankton. Nature 345:60–63. Giraud, E., J. Fardoux, N. Fourrier, L. Hannibal, B. Genty, P. Bouyer, B. Dreyfus, and A. Vermeglio. 2002. Bacterio-

phytochrome controls photosystem synthesis in anoxygenic bacteria. Nature 417:202–205. Goodwin, C. S., J. A. Armstrong, T. Chilvers, M. Peters, M. D. Collins, L. Sly, W. McConnell, and W. E. S. Harper. 1989. Transfer of Campylobacter pylori and Campylobacter mustelae to Helicobacter gen. nov., as Helicobacter pylori comb. nov. and Helicobacter mustelae comb. nov., respectively. Int. J. Syst. Bacteriol. 39:397– 405. Goris, J., P. De Vos, T. Coenye, B. Hoste, D. Janssens, H. Brim, L. Diels, M. Mergeay, K. Kersters, and P. Vandamme. 2001. Classification of metal-resistant bacteria from industrial biotopes as Ralstonia campinensis sp. nov., Ralstonia metallidurans sp. nov.and Ralstonia basilensis Steinle et al. 1998 emend. Int. J. Syst. Evol. Microbiol. 51:1773–1782. Govan, J. R. W., R. Balandreau, and P. Vandamme. 2000. Burkholderia cepacia - friend and foe. ASM News 66: 124–125. Gray, M. W., and W. F. Doolittle. 1982. Has the endosymbiont hypothesis been proven? Microbiol. Rev. 46:1–42. Gray, M. W. 1998. Rickettsia, typhus and the mitochondrial connection. Nature 396:109–110. Gray M. W., G. Burger, and B. F. Lang. 1999. Mitochondrial evolution. Science 283:1476–1481. Gros, O., P. De Wulf-Durand, L. Frenkiel, and M. Moueza. 1998. Putative environmental transmission of sulfuroxidizing bacterial symbionts in tropical lucinid bivalves inhabiting various environments. FEMS Microbiol. Lett. 160:257–262. Gupta, R. S. 2000. The phylogeny of proteobacteria: relationships to other eubacterial phyla and eukaryotes. FEMS Microbiol. Rev. 24:367–402. Gupta, R. S. 2002. Phylogeny of Bacteria: are we now close to understanding it? ASM News 68:284–291. Hamana, K., and S. Matsuzaki. 1993. Polyamine distribution patterns serve as a phenotypic marker in the chemotaxonomy of the proteobacteria. Can. J. Microbiol. 39:304– 310. Hauben, L., E. R. B. Moore, L. Vauterin, M. Steenackers, J. Mergaert, L. Verdonck, and J. Swings. 1998. Phylogenetic position of phytopathogens within the Enterobacteriaceae. Syst. Appl. Microbiol. 21:384–397. Hayashi, N. R., T. Ishida, A. Yokota, T. Kodama, and Y. Igarashi. 1999. Hydrogenophilus thermoluteolus gen, nov., sp. nov., a thermophilic, facultatively chemolithoautotrophic, hydrogen-oxidizing bacterium. Int. J. Syst. Bacteriol. 49:783–786. Haygood, M. G., D. L. Distel, and P. J. Herring. 1992. Polymerase chain reaction and 16S ribosomal RNA gene sequences from the luminous bacterial symbionts of two deep-sea anglerfishes. J. Mar. Biol. Ass. UK 72:149–159. Hedlund, B. P., and J. T. Staley. 2002. Phylogeny of the genus Simonsiella and other members of the Neisseriaceae. Int. J. Syst. Evol. Microbiol. 52:1377–1382. Hippe, H., A. Hagelstein, I., Kramer, J. Swiderski, and E. Stackebrandt. 1999. Phylogenetic analysis of Formivibrio citricus, Propionivibrio dicarboxylicus, Anaerobiospirillum thomasii, Succinimonas amylolytica and Succinivibrio dextrinosolvens and proposal of Succinivibrionaceae fam. nov. Int. J. Syst. Bacteriol. 49:779–782. Hiraishi, A., Y. Hoshino, and H. Kitamura. 1984. Isoprenoid quinone composition in the classification of Rhodospirillaceae. J. Gen. Appl. Microbiol. 30:197–210. Holt, J. G., N. R. Krieg, P. H. A. Sneath, J. T. Staley, and S. T. Williams. 1994. Bergey’s Manual of Determinative Bac-

Introduction to the Proteobacteria teriology, 9th ed. Williams and Wilkins. Baltimore, MD. 1–787. Hookey, J. V., N. A. Saunders, N. K. Fry, R. J. Birtles, and T. G. Harrison. 1996. Phylogeny of Legionellaceae based on small-subunit ribosomal DNA sequences and proposal of Legionella lytica comb. nov. for Legionella-like amoebal pathogens. Int. J. Syst. Bacteriol. 46:526–531. Horn, M., T. R. Fritsche, T. Linner, R. K. Gautom, M. D. Harzenetter, and M. Wagner. 2002. Obligate bacterial endosymbionts of Acanthamoeba spp. related to the beta-proteobacteria: Proposal of “Candidatus Procabacter acanthamoebae” gen. nov., sp. nov. Int. J. Syst. Evol. Microbiol. 52:599–605. Hugenholz, P., P. Goebel, and N. Pace. 1998. Impact of culture-independent studies on the emerging phylogenetic view of bacterial diversity. J. Bacteriol. 180:4765–4774. Hypsa, V., and C. Dale. 1997. In vitro culture and phylogenetic analysis of “Candidatus Arsenophonus triatominarum,” an intracellular bacterium from the triatomine bug, Triatoma infestans. Int. J. Syst. Bacteriol. 47:1140– 1144. Imhoff, J. F., H. G. Trüper, and N. Pfennig. 1984. Rearrangement of the species and genera of the phototrophic “purple nonsulfur bacteria.” Int. J. Syst. Bacteriol. 34:340– 343. Imhoff, J. F., R. Petri, and J. Süling. 1998. Reclassification of species of the spiral-shaped phototrophic purple nonsulfur bacteria of the alpha-proteobacteria: Description of the new genera Phaeospirillum gen. nov., Rhodovibrio gen. nov., Rhodothalassium gen. nov. and Roseospira gen. nov. as well as transfer of Rhodospirillum fulvum to Phaeospirillum fulvum comb. nov., of Rhodospirillum molischianum to Phaeospirillum molischianum comb. nov., of Rhodospirillum salinarum to Rhodovibrio salinarum comb. nov., of Rhodospirillum sodomense to Rhodovibrio sodomensis comb. nov., of Rhodospirillum salexigens to Rhodothalassium salexigens comb. nov. and of Rhodospirillum mediosalinum to Roseospira mediosalina comb. nov. Int. J. Syst. Bacteriol. 48:793–798. Imhoff, J. F. 2001a. The anoxygenic phototrophic purple bacteria. In: G. Garrity (Ed.) Bergey’s Manual of Systematic Bacteriology, 2nd ed. Springer-Verlag. New York, NY. 1:631–637. Imhoff, J. F. 2001b. True marine and halophilic anoxygenic phototrophic bacteria. Arch. Microbiol. 176:243–254. Jacobi, C. A., B. Aßmus, H. Reichenbach, and E. Stackebrandt. 1997. Molecular evidence for association between the Sphingobacterium-like organism “Candidatus comitans” and the myxobacterium Chondromyces crocatus. Appl. Environ. Microbiol. 63:719–723. Jagoueix, S., J. M. Bové, and M. Garnier. 1994. The phloemlimited bacterium of greening disease of citrus is a member of the alpha-subdivision of the Proteobacteria. Int. J. Syst. Bacteriol. 44:379–386. Jarvis, B. D. W., M. Gillis, and J. De Ley. 1986. Intrageneric and intergeneric similarities between the ribosomal ribonucleic acid cistrons of Rhizobium and Bradyrhizobium species and some related bacteria. Int. J. Syst. Bacteriol. 36:129–138. Jumas-Bilak, E. J., S. Michaux-Charachon, G. Bourg, M. Ramuz, and A. Allardet-Servent. 1998. Unconventional genomic organization in the alpha-subgroup of the Proteobacteria. J. Bacteriol. 180:2749–2755. Juretschko, S., A. Loy, A. Lehner, and M. Wagner. 2002. The microbial community composition of a nitrifying-

33

denitrifying activated sludge from an industrial sewage treatment plant analyzed by the full-cycle rRNA approach. System. Appl. Microbiol. 25:84–99. Jurkevitch, E. 2001. The genus Bdellovibrio. In: M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer, and E. Stackebrandt (Eds.) The Prokaryotes, 3rd ed. SpringerVerlag. New York, NY. Karlberg, O., B. Canbäck, C. G. Kurland, and S. G. E. Andersson. 2000. The dual origin of the yeast mitochondrial proteome. Yeast 17:170–187. Kelly, D. P., and A. P. Wood. 2000. Reclassification of some species of Thiobacillus to the newly designated genera Acidithiobacillus gen. nov., Halothiobacillus gen. nov. and Thermithiobacillus gen. nov. Int. J. Syst. Evol. Microbiol. 50:511–516. Kersters, K., and J. De Ley. 1984. Genus Alcaligenes Castellani and Chalmers 1919, 936. In: N. R. Krieg and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology, Vol. 1. Williams and Wilkins. Baltimore, MD. 1:361–373. Kersters, K., W. Ludwig, M. Vancanneyt, P. De Vos, M. Gillis, and K.-H. Schleifer. 1996. Recent changes in the classification of the pseudomonads: An overview. Syst. Appl. Microbiol. 19:465–477. Khan, I. A., and N. J. Mehta. 2002. Stenotrophomonas maltophilia endocarditis: A systematic review. Angiol. 53:49–55. Klein, M., M. Friedrich, A. J. Roger, P. Hugenholtz, S. Fishbain, H. Abicht, L. L. Blackall, D. A. Stahl, and M. Wagner. 2001. Multiple lateral transfers of dissimilatory sulfite reductase genes between major lineages of sulfate-reducing prokaryotes. J. Bacteriol. 183:6028–6035. Klenk, H.-P., T. D. Meier, P. Durovic, V. Schwass, F. Lottspeich, P. P. Dennis, and W. Zillig. 1999. RNA polymerase of Aquifex pyrophilus: implications for the evolution of the bacterial rpoBC operon and extremely thermophilic bacteria. J. Molec. Evol. 48:528–541. Knoblauch, C., K. Sahm, and B. B. Jorgensen. 1999. Psychrophilic sulfate-reducing bacteria isolated from permanently cold Arctic marine sediments: Description of Desulfofrigrus oceanense gen. nov., sp. nov., Desulfofrigus fragile sp. nov., Desulfofaba gelida gen. nov., sp. nov., Desulfotalea psychrophila gen. nov., sp. nov. and Desulfotalea arctica sp. nov. Int. J. Syst. Bacteriol. 49:1631– 1643. Krieg, N. R., and J. G. Holt. 1984. Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 1:964. Krueger, D. M., R. G. Gustafson, and C. M. Cavanaugh. 1996. Vertical transmission of chemoautotrophic symbionts of the bivalve Solemya velum (Bivalvia: Protobranchia). Biol. Bull. 190:195–202. Kurland, C. G., and S. G. E. Andersson. 2000. Origin and evolution of the mitochondrial proteome. Microbiol. Molec. Biol. Rev. 64:786–820. Labrenz, M., M. D. Collins, P. A. Lawson, B. J. Tindall, G. Braker, and P. Hirsch. 1998. Antarctobacter heliothermus gen. nov., sp. nov., a budding bacterium from hypersaline and heliothermal Ekho Lake. Int. J. Syst. Bacteriol. 48:1363–1372. Labrenz, M., B. J. Tindall, P. A. Lawson, M. D. Collins, P. Schumann, and P. Hirsch. 2000. Staleya guttiformis gen. nov., sp. nov. and Sulfitobacter brevis sp. nov., alpha-3proteobacteria from hypersaline, heliothermal and meromictic antarctic Ekho Lake. Int. J. Syst. Evol. Microbiol. 50:303–313.

34

K. Kersters et al.

Lang, B. F., E. Seif, M. W. Gray, C. J. O’Kelly, and G. Burger. 1999. A comparative genomics approach to the evolution of eukaryotes and their mitochondria. J. Eukaryot. Microbiol. 46:320–326. Lee, N., P. H. Nielsen, K. H. Andreasen, S. Juretschko, J. L. Nielsen, K. H. Schleifer, and M. Wagner. 1999. Combination of fluorescent in situ hybridization and microautoradiography: A new tool for structure-function analyses in microbial ecology. Appl. Environ. Microbiol. 65:1289–1297. Lessie, T. G., W. Hendrickson, B. D. Manning, and R. Devereux. 1996. Genomic complexity and plasticity of Burkholderia cepacia. FEMS Microbiol. Lett. 144:117– 128. Liesack, W., and E. Stackebrandt. 1992. Unculturable microbes detected by molecular sequences and probes. Biodivers. Conserv. 1:250–262. Liesack, W., and K. Finster. 1994. Phylogenetic analysis of five strains of Gram-negative, obligately anaerobic, sulfur-reducing bacteria and description of Desulfuromusa gen. nov., including Desulfuromusa kysingii sp. nov., Desulfuromusa bakii sp. nov., and Desulfuromusa succinoxidans sp. nov. Int. J. Syst. Bacteriol. 44:753–758. LiPuma, J. J., T. Spilker, L. H. Gill, P. W. Campbell 3rd, L. X. Liu, and E. Mahenthiralingam. 2001. Disproportionate distribution of Burkholderia cepacia complex species and transmissibility markers in cystic fibrosis. Am. J. Respir. Crit. Care Med. 164:92–96. Ludwig, W., R. Rossellomora, R. Aznar et al. 1995. Comparative sequence analysis of 23S ribosomal-RNA from proteobacteria. Syst. Appl. Microbiol. 18:164–188. Ludwig, W., R. Amann, E. Martinez-Romero, W. Schönhuber, S. Bauer, A. Neef, and K.-H. Schleifer. 1998. rRNA based identification and detection systems for rhizobia and other bacteria. Plant Soil 204:1–19. Ludwig, W., and K.-H. Schleifer. 1999. Phylogeny of bacteria beyond the 16S rRNA standard. ASM News 65:752–757. Ludwig, W., and H.-P. Klenk. 2001. Overview: A phylogenetic backbone and taxonomic framework for procaryotic systematics. In: G. Garrity (Ed.) Bergey’s Manual of Systematic Bacteriology, 2nd ed. Springer-Verlag. Baltimore, MD. 1:49–65. Mahenthiralingam, E., T. Coenye, J. W. Chung, D. P. Speert, J. R. W. Govan, P. Taylor, and P. Vandamme. 2000. Diagnostically and experimentally useful panel of strains from the Burkholderia cepacia complex. J. Clin. Microbiol. 38:910–913. Maiden, M. C. J., J. A. Bygraves, E. Feil, G. Morelli, J. E. Russel, R. Urwin, Q. Zhang, J. Zhou, K. Zurth, D. A. Caugant, I. M. Feavers, M. Achtman, and B. G. Spratt. 1998. Multilocus sequence typing: A portable approach to the identification of clones within populations of pathogenic organisms. Proc. Natl. Acad. Sci. USA 95:3140–3145. Manz, W., R. Amann, W. Ludwig, M. Wagner, and K.-H. Schleifer. 1992. Phylogenetic oligonucleotide probes for the major subclasses of proteobacteria: Problems and solutions. Syst. Appl. Microbiol. 15:593–600. Margulis, L. 1993. Symbiosis in cell evolution: Microbial communities in the archean and proterozoic eons. Freeman. New York, NY. 1–452. Margulis, L. 1996. Archaeal-eubacterial mergers in the origin of Eukarya: Phylogenetic classification of life. Proc. Natl. Acad. Sci. USA 93:1071–1076. Maszenan, A. M., R. J. Seviour, B. K. C. Patel, G. N. Rees, and B. M. McDougall. 1997. Amaricoccus gen. nov., a

Gram-negative coccus occurring in regular packages or tetrads, isolated from activated sludge biomass, and descriptions of Amaricoccus veronensis sp. nov., Amaricoccus tamworthensis sp. nov., Amaricoccus macauensis sp. nov., and Amaricoccus kaplicensis sp. nov. Int. J. Syst. Bacteriol. 47:727–734. Maszenan, A. M., R. J. Seviour, B. K. C. Patel, and P. Schumann. 2002. Quadricoccus autraliensis gen. nov., sp. nov., a beta-proteobacterium from activated sludge biomass. Int. J. Syst. Evol. Microbiol. 52:223–228. McOrist, S., C. J. Gebhart, R. Boid, and S. M. Barns. 1995. Characterization of Lawsonia intracellularis gen. nov., sp. nov., the obligately intracellular bacterium of porcine proliferative enteropathy. Int. J. Syst. Bacteriol. 45:820– 825. McOrist, A. L., M. Warhurst, S. McOrist, and A. R. Bird. 2001. Colonic infection by Bilophila wadsworthia in pigs. J. Clin. Microbiol. 39:1577–1579. Michaux-Charachon, S., G. Bourg, E. Jumas-Bilak, P. Guigue-Talet, D. O’Callaghan, A. Allardet-Servent, and M. Ramuz. 1997. Genome structure and phylogeny in the genus Brucella. J. Bacteriol. 179:3244–3249. Miroshnichenko, M. L., F. A. Rainey, M. Rhode, and E. A. Bonch-Osmolovskaya. 1999. Hippea maritima gen. nov., sp. nov., a new genus of thermophilic, sulfur-reducing bacterium from submarine hot vents. Int. J. Syst. Bacteriol. 49:1033–1038. Miroshnichenko, M. L., N. A. Kostrikina, S. L’Haridon, C. Jeanthon, H. Hippe, E. Stackebrandt, and E. A. Bonch-Osmolovskaya. 2002. Nautilia lithotrophica gen. nov., sp. nov., a thermophilic sulfur-reducing epsilonproteobacterium isolated from a deep-sea hydrothermal vent. Int. J. Syst. Evol. Microbiol. 52:1299–1304. Molouba, F., J. Lorquin, A. Willems, B. Hoste, E. Giraud, B. Dreyfus, M. Gillis, P. de Lajudie, and C. MassonBoivin. 1999. Photosynthetic bradyrhizobia from Aeschynomene spp. are specific to stem-nodulated species and form a separate 16S ribosomal DNA restriction fragment length polymorphism group. Appl. Environ. Microbiol. 65:3084–3094. Moran, N. A., M. A. Munson, P. Baumann, and H. Ishikawa. 1993. A molecular clock in endosymbiotic bacteria is calibrated using the insect host. Proc. Royal Soc. London Ser. B Biol. Sci. 235:167–171. Moss, C. W., M. A. Lambert-Fair, M. A. Nicholson, and G. O. Guerrant. 1990. Isoprenoid quinones of Campylobacter cryaerophila, C. cinaedi, C. fennelliae, C. hyointestinalis, C. pylori, and “C. upsaliensis.” J. Clin. Microbiol. 28:395– 397. Moulin, L., A. Munive, B. Dreyfus, and C. Boivin-Masson. 2001. Nodulation of legumes by members of the betasubclass of proteobacteria. Nature 411:948–950. Munson, M. A., P. Baumann, and N. A. Moran. 1992. Phylogenetic relationships of the endosymbionts of the mealybugs (Homoptera: Pseudococcidae) based on 16S rDNA sequences. Molec. Phylogenet. Evol. 1:26–30. Murray, R. G. E., and K.-H. Schleifer. 1994. Taxonomic notes: a proposal for recording the properties of putative taxa of prokaryotes. Int. J. Syst. Bacteriol. 44:174–176. Murray, R. G. E., and E. Stackebrandt. 1995. Taxonomic note: Implementation of the provisional status Candidatus for incompletely described procaryotes. Int. J. Syst. Bacteriol. 45:186–187. Ochman, H., and A. C. Wilson. 1987. Evolution in bacteria: Evidence for a universal substitution rate in cellular genomes. J. Molec. Evol. 26:74–86.

Introduction to the Proteobacteria Olson, J. M. 1970. The evolution of photosynthesis. Science 168:438–446. On, S. L. W. 2001. Taxonomy of campylobacter, arcobacter, helicobacter and related bacteria: current status, future prospects and immediate concerns. J. Appl. Microbiol. 90:1S–15S. Ouverney, C. C., and J. A. Fuhrman. 1999. Combined microautoradiography-16S rRNA probe technique for determination of radioisotope uptake by specific microbial cell types in situ. Appl. Environ. Microbiol. 65:1746– 1752. Oyaizu, H., and C. R. Woese. 1985. Phylogenetic relationships among the sulfate respiring bacteria, myxobacteria and purple bacteria. Syst. Appl. Microbiol. 6:257–263. Palleroni, N. J. 1984. Pseudomonas Migula 1894, 237. In: N. R. Krieg and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 1:141–199. Parke, J. L., and D. Gurian-Sherman. 2001. Diversity of the Burkholderia cepacia complex and implications for risk assessment of biological control strains. Ann. Rev. Phytopathol. 39:225–258. Parkhill, J., B. W. Wren, K. Mungall et al. 2000. The genome sequence of the food-borne pathogen Campylobacter jejuni reveals hypervariable sequences. Nature 403:665– 668. Rabus, R., M. Fukui, H. Wilkes, and F. Widdel. 1996. Degradative capacities and 16S rDNA-targeted whole cell hybridization of sulfate-reducing bacteria in an anaerobic enrichment culture utilizing alkylbenzenes from crude oil. Appl. Environ. Microbiol. 62:3605–3613. Rahn, O. 1937. New principles for the classification of bacteria. Zentralbl. Bakteriol. Parasitenkd. Infectionskr. Hyg., Abt. II 96:273–286. Rainey, F. A., R. Toalster, and E. Stackebrandt. 1993. Desulfurella acetivorans, a thermophilic, acetate-oxidizing and sulfur-reducing organism, represents a distinct lineage within the proteobacteria. Syst. Appl. Microbiol. 16:373–379. Reichenbach, H., and G. Hofle. 1993. Biologically-active secondary metabolites from myxobacteria. Biotechnol. Adv. 11:219–277. Reichenbach, H. 2001. Myxobacteria, producers of novel bioactive substances. J. Ind. Microbiol. Biotechnol. 27: 149–156. Romanenko, L. A., P. Schumann, M. Rohde, V. V. Mikhailov, and E. Stackebrandt. 2002. Halomonas halocynthiae sp. nov., a novel bacterium, isolated from the marine ascidian Halocynthia aurantium. Int. J. Syst. Evol. Microbiol. [doi 10.1099/ijs.0.02240-0] IJSEM Papers in Press. 52:1767–1772. Romaniuk, P. J., B. Zoltowska, T. J. Trust, D. J. Lane, G. J. Olsen, N. R. Pace, and D. A. Stahl. 1987. Campylobacter pylori, the spiral bacterium associated with human gastritis, is not a true campylobacter sp. J. Bacteriol. 169:2137–2141. Rossau, R., A. Van Landschoot, W. Mannheim, and J. De Ley. 1986. Intergeneric and intrageneric similarities of ribosomal ribonucleic acid cistrons of the Neisseriaceae. Int. J. Syst. Bacteriol. 36:323–332. Rossau, R., A. Van Landschoot, M. Gillis, and J. De Ley. 1991. Taxonomy of Moraxellaceae fam. nov., a new bacterial family to accommodate the genera Moraxella, Acinetobacter, and Psychrobacter and related organisms. Int. J. Syst. Bacteriol. 41:310–319.

35

Rossellomora, R. A., W. Ludwig, and K.-H. Schleifer. 1993. Zoogloea ramigera: A phylogenetically diverse species. FEMS Microbiol. Lett. 114:129–134. Sahm H., S. Bringer-Meyer, and G. A. Sprenger. 2001. The genus Zymomonas. In: M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer, and E. Stackebrandt (Eds.) The Prokaryotes, 3rd ed. Springer-Verlag. New York, NY. Satomi, M., B. Kimura, T. Hamada, S. Harayama, and T. Fujii. 2002. Phylogenetic study of the genus Oceanospirillum based on 16S rRNA and gyrB genes: emended description of the genus Oceanospirillum, description of Pseudospirillum gen. nov., Oceanobacter gen. nov. and Terasakiella gen. nov. and transfer of Oceanospirillum jannaschii and Pseudomonas stanieri to Marinobacterium as Marinobacterium jannaschii comb. nov. and Marinobacterium stanieri comb. nov. Int. J. Syst. Evol. Microbiol. 52:739–747. Sauer, C., E. Stackebrandt, J. Gadau, B. Hölldobler, and R. Gross. 2000. Systematic relationships and cospeciation of bacterial endosymbionts and their carpenter ant host species: proposal of the new taxon Candidatus Blochmannia gen. nov. Int. J. Syst. Evol. Microbiol. 50: 1877–1886. Scholz-Muramatsu, H., A. Neumann, M. Messmer, E. Moore, and G. Diekert. 1995. Isolation and characterization of Dehalospirillum multivorans gen. nov. sp. nov., a tetrachloroethene utilizing, strictly anaerobic bacterium. Arch. Microbiol. 163:48–56. Schröder, D., H. Deppisch, M. Obermeyer, G. Krohne, E. Stackebrandt, B. Hoelldobler, W. Göbel, and R. Gross. 1996. Intracellular endosymbiotic bacteria of Camponotus species (carpenter ants): systematics, evolution and ultrastructural characterization. Molec. Microbiol. 21: 479–489. Schulz, H. N., T. Brinkhoff, T. G. Ferdelman, M. H. Marine, A. Teske, B. B. Jorgensen. 1999. Dense populations of a giant sulfur bacterium in Namibian shelf sediments. Science 284:493–495. Schulz, H. N. 2002. Thiomargarita namibiensis: giant microbe holding its breath. ASM News 68:122–127. Schwartz, R. M., and M. O. Dayhoff. 1978. Origins of prokaryotes, eukaryotes, mitochondria and chloroplasts. Science 199:395–403. Schwemmler, W. 1989. Symbiogenesis: A macro-mechanism of evolution. Walter de Gruyter. Berlin, Germany. 1–226. Seewaldt, E., K.-H. Schleifer, E. Bock, and E. Stackebrandt. 1982. The close phylogenetic relationship of Nitrobacter and Rhodopseudomonas palustris. Arch. Microbiol. 131:287–290. Seidler, R. J., M. Mandel, and J. N. Baptist. 1972. Molecular heterogeneity of Bdellovibrio: Evidence of two new species. J. Bacteriol. 109:207–217. Simpson, A. J. G., F. C. Reinach, P. Arruda et al. 2000. The genome sequence of the plant pathogen Xylella fastidiosa. Nature 406:151–157. Snaidr, J., R. Amann, I. Huber, W. Ludwig, and K.-H. Schleifer. 1997. Phylogenetic analysis and in situ identification of bacteria in activated sludge. Appl. Environ. Microbiol. 63:2884–2896. Sobral, B. W. S., R. J. Honeycutt, A. G. Atherly, and M. McClelland. 1991. Electrophoretic separation of the three Rhizobium meliloti replicons. J. Bacteriol. 173: 5173–5180. Solnick, J. V., and P. Vandamme. 2001. Taxonomy of the Helicobacter genus. In: H. L. T. Mobley, G. L. Mendz, and

36

K. Kersters et al.

S. L. Hazell (Eds.) Helicobacter pylori Physiology and Genetics. ASM Press. Washington, DC. 39–51. Sorokin, D. Y., A. M. Lysenko, L. L. Mityushina, T. P. Tourova, B. E. Jones, F. A. Rainey, L. A. Robertson, and G. J. Kuenen. 2001. Thioalkalimicrobium aerophilum gen. nov., sp. nov. and Thioalkalimicrobium sibericum sp. nov., and Thioalkalivibrio versutus gen. nov., sp. nov., Thioalkalivibrio nitratis sp. nov. and Thioalkalivibrio denitrificans sp. nov., novel obligately alkaliphilic and obligately chemolithoautotrophic sulfur-oxidizing bacteria from soda lakes. Int. J. Syst. Evol. Microbiol. 51:565–580. Spormann, A. M. 1999. Gliding motility in bacteria: insights from studies of Myxococcus xanthus. Microbiol. Molec. Biol. Rev. 63:621–641. Spring, S., P. Kämpfer, W. Ludwig, and K.-H. Schleifer. 1996. Polyphasic characterization of the genus Leptothrix: new descriptions of Leptothrix mobilis sp. nov. and Leptothrix discophora sp. nov. nom. rev. and emended description of Leptothrix cholodnii emend. Syst. Appl. Microbiol. 19:634–643. Spröer, C., H. Reichenbach, and E. Stackebrandt. 1999. The correlation between morphological and phylogenetic classification of myxobacteria. Int. J. Syst. Bacteriol. 49:1255–1262. Stackebrandt, E., T. M. Embley, and J. Weckesser. 1988a. Phylogenetic, evolutionary and taxonomic aspects of phototrophic eubacteria. In: J. M. Olson, J. G. Ormerod, J. Amesz, E. Stackebrandt, and H. G. Trüper (Eds.) Green Phototrophic Bacteria. Plenum Press. New York, NY. 201–215. Stackebrandt, E., R. G. E. Murray, and H. G. Trüper. 1988b. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “purple bacteria and their relatives.” Int. J. Syst. Bacteriol. 38:321–325. Stackebrandt, E. 1992. Unifying phylogeny and phenotypic diversity. In: A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York, NY. 19–47. Stackebrandt, E. 1995. Bacterial phylogeny. In: A. J. Gibbs, C. H. Calisher, and F. Garcia-Arenal (Eds.) Molecular Basis of Virus Evolution. Academic Press. London, UK. 15–28. Stackebrandt, E., F. A. Rainey, and N. Ward-Rainey. 1996. Anoxygenic phototrophy across the phylogenetic spectrum: current understanding and future perspectives. Arch. Microbiol. 166:211–223. Stackebrandt, E., W. Frederiksen, G. M. Garrity, P. A. D. Grimont., P. Kämpfer, M. C. J. Maiden, X. Nesme, R. Rosselló-Mora, J. Swings, H. G. Trüper, L. Vauterin, A. C. Ward, and W. B. Whitman. 2002. Report of the ad hoc committee for the re-evaluation of the species definition in bacteriology. Int. J. Syst. Evol. Microbiol 52: 1043–1047. Stahl, D. A., and M. Wagner. 2001. Multiple lateral transfer events of dissimilatory sulfite reductase genes between major lineages of sulfate-reducing prokaryotes. J. Bacteriol. 183:6028–6035. Staley, J. T., and A. Konopka. 1985. Measurement of in situ activities of nonphotosynthetic microorganisms in aquatic and terrestrial habitats. Ann. Rev. Microbiol. 39:321–346. Stolz, J. F., D. J. Ellis, J. S. Blum, D. Ahmann, D. R. Lovley, and R. S. Oremland. 1999. Sulfurospirillum barnesii sp nov and Sulfurospirillum arsenophilum sp nov., new members of the Sulfurospirillum clade of the epsilon proteobacteria. Int. J. Syst. Bacteriol. 49:1177–1180.

Stouthamer, R., J. A. J. Breeuwer, R. F. Luck, and J. H. Werren. 1993. Molecular identification of microorganisms associated with parthenogenesis. Nature 361:66–68. Suwanto, A., and S. Kaplan. 1989. Physical and genetic mapping of the Rhodobacter sphaeroides 2.4.1 genome: Presence of two unique circular chromosomes. J. Bacteriol. 171:5850–5859. Suyama, T, T. Shigematsu, S. Takaichi, Y. Nodasaka, S. Fujikawa, H. Hosoya, Y. Tokiwa, T. Kanagawa, and S. Hanada. 1999. Roseateles depolymerans gen, nov., sp. nov., a new bacteriochlorophyll a-containing obligate aerobe belonging to the beta-subclass of the proteobacteria. Int. J. Syst. Bacteriol. 49:449–457. Swings, J., and J. De Ley. 1977. Biology of Zymomonas. Bacteriol. Rev. 41:1–46. Swings, J. 1992. The genera Acetobacter and Gluconobacter. In: A. Balows A., H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York , NY. 2268–2286. Sy, A., E. Giraud, P. Jourand, N. Garcia, A. Willems, P. de Lajudie, Y. Prin, M. Neyra, M. Gillis, C. Boivin-Masson, and B. Dreyfus. 2001. Methylotrophic Methylobacterium bacteria nodulate and fix nitrogen in symbiosis with legumes. J. Bacteriol. 183:214–220. Takeuchi, M., K. Hamana, and A. Hiraishi. 2001. Proposal of the genus Sphingomonas sensu stricto and three new genera, Sphingobium, Novosphingobium and Sphingopyxis, on the basis of phylogenetic and chemotaxonomic analyses. Int. J. Syst. Evol. Microbiol. 51:1405–1417. Tamaoka, J., D. M. Ha, and K. Komagata. 1987. Reclassification of Pseudomonas acidovorans Den Dooren De Jong 1926 and Pseudomonas testosteroni Marcus and Talalay 1956 as Comamonas acidovorans comb. nov. and Comamonas testosteroni comb. nov., with an emended description of the genus Comamonas. Int. J. Syst. Evol. Bacteriol. 37:52–59. Tanaka, K., E. Stackebrandt, S. Tohyama, and T. Eguchi. 2000. Desulfovirga adipica gen. nov., sp. nov., an adipatedegrading, Gram-negative, sulfate-reducing bacterium. Int. J. Syst. Evol. Bacteriol. 50:639–644. Teske, A., P. Sigalevich, Y. Cohen, and G. Muyzer. 1996. Molecular identification of bacteria from a coculture by denaturing gradient gel electrophoresis of 16S ribosomal DNA fragments as a tool for isolation in pure cultures. Appl. Environ. Microbiol. 62:4210–4215. Unterman, B. M., P. Baumann, and D. L. McLean. 1989. Pea aphid symbiont relationships established by analysis of 16S rRNAs. J. Bacteriol. 171:2970–2974. Urakawa, H., T. Yoshida, M. Nishimura, and K. Ohwada. 2000. Characterization of depth-related population variation in microbial communities of a coastal marine sediment using 16S rDNA-based approaches and quinone profiling. Environ. Microbiol. 2:542–554. Van Couwenberghe, C. J., T. B. Farver, and S. H. Cohen. 1997. Risk factors associated with isolation of Stenotrophomonas (Xanthomonas) maltophilia in clinical specimens. Infect. Cont. Hosp. Epidemiol. 18:316–321. Vandamme, P., and J. De Ley. 1991. Proposal for a new family, Campylobacteraceae. Int. J. Syst. Bacteriol. 41:451–455. Vandamme, P., B. Pot, M. Gillis, P. De Vos, K. Kersters, and J. Swings. 1996. Polyphasic taxonomy, a consensus approach to bacterial systematics. Microbiol. Rev. 60:407–438. Vandamme, P., B. Holmes, M. Vancanneyt, T. Coenye, B. Hoste, R. Coopman, H. Revets, S. Lauwers, M. Gillis, K. Kersters, and J. R. W. Govan. 1997. Occurrence of mul-

Introduction to the Proteobacteria tiple genomovars of Burkholderia cepacia in cystic fibrosis patients and proposal of Burkholderia multivorans sp. nov. Int. J. Syst. Bacteriol. 47:1188–1200. Vandamme, P., P. Segers, M. Ryll, J. Hommez, M. Vancanneyt, R. Coopman, R. De Baere, Y. Van de Peer, K. Kersters, and R. De Wachter. 1998. Pelistega europaea gen. nov., sp. nov., a bacterium associated with respiratory disease in pigeons: taxonomic structure and phylogenetic allocation. Int. J. Syst. Bacteriol. 48:431–440. Vandamme, P. 2000. Taxonomy of the family Campylobacteraceae. In: I. Nachamkin and M. Blaser (Eds.) Campylobacter (2nd ed.). ASM Press. Washington, DC. 3–26. Vandamme, P., and S. L. W. On. 2001. Recommendations of the Subcommittee on the taxonomy of Campylobacter and related bacteria. Int. J. Syst. Evol. Microbiol. 51:719–721. Van de Peer, Y., and R. De Wachter. 1997. Construction of evolutionary distance trees with TREECON for Windows: accounting for variation in nucleotide substitution rate among sites. Comput. Appl. Biosci. 13:227–230. Voordouw, G., S. M. Armstrong, M. F. Reimer, B. Fouts, A. J. Telang, Y. Shen, and D. Gevertz. 1996. Characterization of 16S rRNA genes from oil field microbial communities indicates the presence of a variety of sulfate-reducing, fermentative, and sulfide-oxidizing bacteria. Appl. Environ. Microbiol. 62:1623–1629. Wagner, M., R. Amann, H. Lemmer, and K.-H. Schleifer. 1993. Probing activated sludge with oligonucleotides specific for proteobacteria: Inadequacy of culturedependent methods for describing microbial community structure. Appl. Environ. Microbiol. 59:1520–1525. Wassenaar, T. M., and D. G. Newell. 2001. The genus Campylobacter. In: M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer, and E. Stackebrandt (Eds.) The Prokaryotes, 3rd ed. Springer-Verlag. New York, NY. Willems, A., J. De Ley, M. Gillis, and K. Kersters. 1991a. Comamonadaceae, a new family encompassing the acidovorans rRNA complex, including Variovorax paradoxus gen. nov., comb. nov., for Alcaligenes paradoxus (Davis 1969). Int. J. Syst. Bacteriol. 41:445–450. Willems, A., M. Gillis, and J. De Ley. 1991b. Transfer of Rhodocyclus gelatinosus to Rubrivivax gelatinosus gen. nov., comb. nov., and phylogenetic relationships with Leptothrix, Sphaerotilus natans, Pseudomonas saccharophila, and Alcaligenes latus. Int. J. Syst. Bacteriol. 41:65–71. Willems, A., H. Gilhaus, W. Beer, H. Mietke, H. R. Gelderblom, B. Burghardt, W. Voigt, and R. Reissbrodt. 2002. Brackiella oedipodis gen. nov., sp nov., Gramnegative, oxidase-positive rods that cause endocarditis of cotton-topped tamarin (Saguinus oedipus). Int. J. Syst. Evol. Microbiol. 52:179–186. Wirsen, C. O., S. M. Sievert, C. M. Cavanaugh, S. J. Molyneaux, A. Ahmad, L. T. Taylor, E. F. DeLong, and C. D. Taylor. 2002. Characterization of an autotrophic sulfide-oxidizing marine Arcobacter sp. that produces filamentous sulfur. Appl. Environ. Microbiol. 68:316– 325. Woese, C. R., and G. E. Fox. 1977. Phylogenetic structure of the prokaryotic domain: the primary kingdoms. Proc. Natl. Acad. Sci. 74:5088–5090.

37

Woese, C. R., E. Stackebrandt, W. G. Weisburg, B. J. Paster, M. T. Madigan, V. J. Fowler, C. M. Hahn, P. Blanz, R. Gupta, K. H. Nealson, and G. E. Fox. 1984a. The phylogeny of purple bacteria: the alpha subdivision. Syst. Appl. Microbiol. 5:315–326. Woese, C. R., W. G. Weisburg, B. J. Paster, C. M. Hahn, R. S. Tanner, N. R. Krieg, H.-P. Koops, H. Harms, and E. Stackebrandt. 1984b. The phylogeny of purple bacteria: The beta-division. Syst. Appl. Microbiol. 5:327– 336. Woese, C. R., B. A. Debrunner-Vossbrinck, H. Oyaizu, E. Stackebrandt, and W. Ludwig. 1985a. Gram-positive bacteria: Possible photosynthetic ancestry. Science 229:762–765. Woese, C. R., E. Stackebrandt, T. J. Macke, and G. E. Fox. 1985b. A phylogenetic definition of the major eubacterial taxa. Syst. Appl. Microbiol. 6:143–151. Woese, C. R., W. G. Weisburg, C. M. Hahn, B. J. Paster, L. B. Zablen, B. J. Lewis, T. J. Macke, W. Ludwig, and E. Stackebrandt. 1985c. The phylogeny of purple bacteria: The gamma subdivision. Syst. Appl. Microbiol. 6:25–33. Woese, C. R. 1987. Bacterial evolution. Microbiol. Rev. 51:221–271. Yabuuchi, E., Y. Kosako, I. Yano, H. Hotta, and Y. Nishiuchi. 1995. Transfer of two Burkholderia and an Alcaligenes species to Ralstonia gen. nov.: Proposal of Ralstonia pickettii (Ralston, Palleroni and Doudoroff 1973) comb. nov., Ralstonia solanacearum (Smith 1896) comb. nov. and Ralstonia eutropha (Davis 1969) comb. nov. Microbiol. Immunol. 39:897–904. Yakimov, M. M., L. Giuliano, T. N. Chernikova, G. Gentile, W.-R. Abraham, H. Lünsdorf, K. N. Timmis, and P. N. Golyshin. 2001. Alcalilimnicola halodurans gen.nov., sp. nov., an alkaliphilic, moderately halophilic and extremely halotolerant bacterium, isolated from sediments of soda-depositing Lake Natron, East Africa Rift Valley. Int. J. Syst. Evol. Microbiol. 51:2133–2143. Young, J. M., L. D. Kuykendall, E. Martinez-Romero, A. Kerr, and H. Sawada. 2001. A revision of Rhizobium Frank 1889, with an emended description of the genus, and the inclusion of all species of Agrobacterium Conn 1942 and Allorhizobium undicola de Lajudie et al. 1998 as new combinations: Rhizobium radiobacter, R. rhizogenes, R. rubi, R. undicola and R. vitis. Int. J. Syst. Evol. Microbiol. 51:89–103. Yurkov, V. V., and J. T. Beatty. 1998. Aerobic anoxygenic phototrophic bacteria. Microbiol. Molec. Biol. Rev. 62:695–724. Yurkov V. V. 2001. Aerobic phototrophic bacteria. In: M. Dworkin, S. Falkow, E. Rosenberg, K.-H. Schleifer, and E. Stackebrandt (Eds.) The Prokaryotes, 3rd ed. Springer-Verlag. New York, NY. Zreik, L., J. M. Bové, and M. Garnier. 1998. Phylogenetic characterization of the bacterium-like organism associated with marginal chlorosis of strawberry and proposition of a Candidatus taxon for the organism, “Candidatus Phlomobacter fragariae.” Int. J. Syst. Bacteriol. 48:257–261. Zuckerkandl, E., and L. Pauling. 1965. Molecules as documents of evolutionary history. J. Theor. Biol. 8:357– 366.

SECTION 3.1

Alpha Subclass

Prokaryotes (2006) 5:41–64 DOI: 10.1007/0-387-30745-1_2

CHAPTER 3.1.1 ehT

c i hpor totohP

a i re t caboe torP- ahp lA

The Phototrophic Alpha-Proteobacteria JOHANNES F. IMHOFF

Introduction The phototrophic purple α-Proteobacteria are purple nonsulfur bacteria able to perform anoxygenic photosynthesis. Owing to the presence of photosynthetic pigments, cell suspensions appear in various colors from beige, olive-green, peachbrown, brown, brown-red, red or pink and have characteristic absorption spectra. Photosynthetic pigments (bacteriochlorophyll a or b [esterified with phytol or geranylgeraniol] and various types of carotenoids) are located in the cytoplasmic membrane and internal membrane systems (vesicles, lamellae or membrane stacks). Typically, phototrophic α-Proteobacteria grow under anoxic conditions in the light and their phototrophic growth, photosynthetic pigment synthesis and internal membrane formation are inhibited by oxygen but become derepressed at low oxygen tensions. Their metabolism is highly diverse and flexible. The preferred mode of growth is photoorganoheterotrophically, but many species also can grow photolithoautotrophically with molecular hydrogen, sulfide or thiosulfate as photosynthetic electron donor; some also can use ferrous iron. Growth factors are generally required, most commonly biotin, thiamine, niacin and p-aminobenzoic acid; growth of most species is enhanced by small amounts of yeast extract, and some have a complex nutrient requirement. Chemotrophic growth under microoxic to oxic conditions in the dark is common to most of these bacteria; some of them are very sensitive to minor levels of oxygen, while others grow equally well aerobically in the dark at the full atmospheric oxygen tension. Anaerobic dark growth by fermentation and oxidant-dependent growth also may occur.

Phylogeny Phototrophic purple nonsulfur bacteria are a highly diverse and heterogeneous group of bacteria, both phenotypically and genetically. On the basis of 16S rDNA sequence similarities,

phototrophic purple bacteria belong to the α-, βand γ-Proteobacteria (Woese et al., 1984a; Woese et al., 1984b; Woese et al., 1985; Stackebrandt et al., 1988; Woese, 1987). While purple sulfur bacteria are γ-Proteobacteria, purple nonsulfur bacteria are found in the β- and α-Proteobacteria. The phototrophic purple β-Proteobacteria (including Rhodocyclus and relatives) are treated elsewhere in this volume (Imhoff). This chapter deals with the phototrophic purple α-Proteobacteria. Three major phylogenetically distinct groups of phototrophic α-Proteobacteria are recognized (see Figs. 1–3). They are represented by Rhodospirillum and relatives (also called “α-1 Proteobacteria”; Fig. 1), by Rhodopseudomonas and relatives (also called “α-2 Proteobacteria”; Fig. 2) and by Rhodobacter and relatives (also called “α-3 Proteobacteria”; Fig. 3). While species of the α-3 group form a tight phylogenetic cluster, in the α-2 group Rhodomicrobium and Rhodobium species are at a greater distance to the other phototrophic species, and in the α-1 group, Rhodovibrio, Rhodopila and Rhodothalassium species form distinct lines separate from the cluster around Rhodospirillum and Phaeospirillum species. Close relatives of Rhodovibrio species and Rhodothalassium salexigens are not known and their assignment to the Rhodospirillum group is arbitrary. On the basis of 16S rDNA sequence similarities, chemotaxonomic characteristics and other properties, many representatives of the phototrophic α-Proteobacteria are very closely related to nonphototrophic, strictly chemotrophic bacteria. These similarities are taken as strong indication for the development of many nonphototrophic bacteria from phototrophic ancestors. A few examples of these relations are the following: The acidophilic Rhodopila globiformis is closely related to other acidophilic bacteria including Acidiphilium species (Sievers et al., 1994). Phaeospirillum species demonstrate a close relationship to Magnetospirillum magnetotacticum (Burgess et al., 1993).

42

J.F. Imhoff

CHAPTER 3.1.1 Escherichia coli Halorhodospira halophila DSM 244 Chromatium okenii DSM 169T Rhodothalassium salexigens ATCC 35888T Rhododovibrio salinarum ATCC 35394T Rhododovibrio sodomense ATCC 51195T Rhodopila globiformis DSM 161T

Fig. 1. Phylogenetic tree based on 16S rDNA sequences of the Rhodospirillum-group (α-1 Proteobacteria) in relation to other purple nonsulfur and purple sulfur bacteria and to representative close chemotrophic relatives.

Acidiphilium angustum ATCC 35903 Acidiphilium cryptum ATCC 33463 Roseospirillum parvum 9301 Rhodospirillum photometricum DSM 122T Rhodospirillum rubrum ATCC 11170T Rhodospira trueperi ATCC 700224T Roseaspira mediosalina BN 280T Magnetospirillum magnetotacticum Phaeospirillum fulvum DSM 113T Phaeospirillum molischianum ATCC 14031T Azospirillum amazonense DSM 2787 Azospirillum halopraeferens DSM 3675 Azospirillum brasilense DSM 2298 Azospirillum irakense 103312 Rhodocista centenaria ATCC 43720T 0.1

Escherichia coli Chromatium okenii DSM 169T Halorhodospira halophila DSM 244T Rhodospirillum rubrum ATCC 11170T Phaeospirillum fulvum DSM 113T Rhodomicrobium vannielii Rhodoblastus acidophilus DSM 137T Rhodopseudomonas palustris ATCC 17001T Nitrobacter winogradskyi ATCC 25381 Nitrobacter hamburgensis K14 Rhodoplanes roseus DSM 5909T Rhodoplanes elegans ATCC 51906T Blastochloris viridis ATCC 19567T Blastochloris sulfoviridis DSM 729T Rhodobium marinum DSM 2698T Rhodobium orientis ATCC 51972T 0.1

Rhodocista centenaria has strong relations to Azospirillum species (Xia et al., 1994; Fani et al., 1995). Within the α-2 Proteobacteria, Rhodopseudomonas palustris has strong relations to species of Nitrobacter (Seewaldt et al., 1982). Rhodobacter and Rhodovulum species form a cluster closely associated to Paracoccus denitrificans (Imhoff, 1989a; Hiraishi and Ueda, 1994a).

Fig. 2. Phylogenetic tree based on 16S rDNA sequences of the Rhodopseudomonas-group (α-2 Proteobacteria) in relation to other purple nonsulfur and purple sulfur bacteria and to representative close chemotrophic relatives.

Taxonomy Since Molisch removed the purple sulfur bacteria from the Thiobacteria (Migula, 1900), pigmentation and ability to perform anoxygenic photosynthesis were considered of primary importance for assignment of bacteria to the Rhodobacteria (Molisch, 1907), later called “Rhodospirillales” (Pfennig and Trüper, 1971).

CHAPTER 3.1.1 Fig. 3. Phylogenetic tree based on 16S rDNA sequences of the Rhodobacter-group (α-3 Proteobacteria) in relation to other purple nonsulfur and purple sulfur bacteria and to representative close chemotrophic relatives.

The Phototrophic Alpha-Proteobacteria

43

Escherichia coli Phaeospirillum fulvum DSM 113T Rhodospirillum rubrum ATCC 11170T Rhodobium marinum ATCC 35601T Rhodopseudomonas palustris ATCC 17001T Blastochloris sulfoviridis DSM 729T Blastochloris viridis ATCC 19567T Rhodovulum euryhalinum DSM 4868T Rhodovulum strictum JCM 9220T Rhodovulum sulfidophilum DSM 1374T Rhodovulum iodosum DSM 12328T Rhodovulum adriaticum DSM 2781T Rhodovulum robiginosum DSM 12329T Rhodobaca bogoriensis LBB1 Roseobacter denitrificans Och 114 Paracoccus denitrificans ATCC 17741T Paracoccus versutus ATCC 25364 Rhodobacter veldkampii ATCC 35703T Rhodobacter blasticus ATCC 33485T Rhodobacter capsulatus ATCC 11166T Rhodobacter azotoformans JCM 9340T Rhodobacter sphaeroides ATCC 17023T

0.1

Because the Rhodospirillaceae (Pfennig and Trüper, 1971) do not represent a phylogenetically distinct group of bacteria, the use of the term “purple nonsulfur bacteria” (PNSB) was proposed for the α- and β-Proteobacteria that contain photosynthetic pigments and are able to perform anoxygenic photosynthesis under anoxic conditions (Imhoff et al., 1984b; Imhoff and Trüper, 1989b; Imhoff and Trüper, 1992b). Actual and historical aspects of the taxonomy of anoxygenic phototrophic purple bacteria have been discussed elsewhere (Imhoff, 1992a; Imhoff, 1995a; Imhoff, 1999; Imhoff, 2000). Traditionally, purple nonsulfur bacteria have been classified into genera representing the rodshaped Rhodopseudomonas species and the spiral-shaped Rhodospirillum species (Pfennig and Trüper, 1974b) and later into a third genus containing the half-circle- to circle-shaped Rhodocyclus purpureus (Pfennig, 1978). With the recognition of their genetic relationships and chemotaxonomic diversity, purple nonsulfur bacteria of the α- and β-Proteobacteria were taxonomically separated (Imhoff et al., 1984b; Imhoff and Trüper, 1989b). Later, bacteria within these groups were rearranged according to phylogeny, chemotaxonomic characteristics and ecophysiological properties. Despite the fact that many of the phototrophic purple nonsulfur bacteria are closely related to strictly chemotrophic relatives, the genus definitions of genera of the anoxygenic

phototrophic bacteria still include phototrophic capability and content of photosynthetic pigments. At higher taxonomic ranks, phototrophic bacteria are treated together with nonphototrophic relatives.

Alpha-1 Proteobacteria Most of the phototrophic bacteria that belong to the α-1 Proteobacteria (also known as the Rhodospirillum group) have been previously known as Rhodospirillum species and are of spiral shape. At present, the only nonspiral representative is Rhodopila globiformis. Genera included in this group are Rhodospirillum, Phaeospirillum, Rhodospira, Roseospira, Rhodocista, Roseospirillum and also Rhodopila, Rhodothalassium and Rhodovibrio. In addition, 16S rDNA sequence data of purple nonsulfur bacteria implied that the spiralshaped phototrophic α-Proteobacteria are phylogenetically quite distantly related to each other and do not warrant classification in one and the same genus (Kawasaki et al., 1993a; Imhoff et al., 1998). These bacteria also demonstrate great phenotypic diversity. Therefore, a reclassification of the spiral-shaped phototrophic α-Proteobacteria was proposed, based on distinct phenotypic properties and 16S rDNA sequence similarities. Rhodospirillum centenum was transferred to a new genus as Rhodocista

44

J.F. Imhoff

centenaria (Kawasaki et al., 1992). Other Rhodospirillum species were transferred to the new genera Phaeospirillum, Rhodovibrio, Roseospira and Rhodothalassium (Imhoff et al., 1998). Only R. rubrum and R. photometricum were maintained as species of the genus Rhodospirillum. In addition, new species were described of this group: Rhodospira trueperi was assigned to a new genus on the basis of significant phenotypic and genotypic differences from Rhodospirillum rubrum and other known PNSB (Pfennig et al., 1997); for similar reasons, the new bacterium Roseospirillum parvum was assigned to a new genus (Glaeser and Overmann, 1999).

Alpha-2 Proteobacteria Most characteristic of phototrophic bacteria of the α-2 (Rhodopseudomonas) group is the budding mode of growth and cell division and the presence of lamellar internal membranes lying parallel to the cytoplasmic membrane. Most of these phototrophic bacteria have been previously known as Rhodopseudomonas species. Genera of this group now include Rhodopseudomonas, Rhodoplanes, Rhodoblastus, Blastochloris, Rhodomicrobium and Rhodobium. After the removal of purple nonsulfur bacteria that contained vesicular internal photosynthetic membranes and those that were β-Proteobacteria from the genus Rhodopseudomonas, only those species remained within this genus that had lamellar internal membrane structures and grew and reproduced by budding (Imhoff et al., 1984b). The bacteria removed from Rhodopseudomonas are now recognized as species of Rhodopila, Rhodobacter, Rhodovulum and Rubrivivax. Thereafter, what remained of the genus Rhodopseudomonas (together with Rhodomicrobium vannielii) still represented a heterogeneous assemblage of species (Imhoff et al., 1984b) now recognized as genera of the α-2 Proteobacteria. Primarily due to the availability of sequence data of the 16S rDNA (Kawasaki et al., 1993a) and in part supported by the isolation and description of new species and additional data, the following proposals have been made. Rhodopseudomonas marina was transferred to the new genus Rhodobium as Rhodobium marinum together with the new species Rhodobium orientis (defined as the type species of this genus; Hiraishi et al., 1995b). Rhodopseudomonas rosea was transferred to the new genus Rhodoplanes and designated as the type species of this genus, R. roseus (Hiraishi and Ueda, 1994b). At the same time, Rhodoplanes elegans was described as a new species of this genus. Rhodopseudomonas viridis and Rhodopseudomonas sulfoviridis were assigned

CHAPTER 3.1.1

to the new genus Blastochloris as B. viridis and B. sulfoviridis (Hiraishi, 1997). Quite recently, Rhodopseudomonas acidophila was transferred to a new genus as Rhodoblastus acidophilus (Imhoff, 2001). Rhodopseudomonas blastica was removed from this genus and transferred to Rhodobacter blasticus (Kawasaki et al., 1993b). Its 16S rDNA sequence is most similar to and clusters with those of the Rhodobacter species. Rhodopseudomonas rutila (Akiba et al., 1983) was considered as a later subjective synonym of Rhodopseudomonas palustris (Hiraishi et al., 1992). In addition to Rhodopseudomonas palustris, Rhodopseudomonas julia (Kompantseva, 1989) and Rhodopseudomonas cryptolactis (Stadtwald-Demchick et al., 1990) have been affiliated to this genus, though both species so far have not been validated and no 16S rDNA sequence of them is available.

Alpha-3 Proteobacteria A characteristic feature of the phototrophic α-3 Proteobacteria (Rhodobacter group) is the presence of carotenoids of the spheroidene series and their extraordinary metabolic versatility and flexibility. These bacteria have been previously known as Rhodopseudomonas species and belong to the genera Rhodobacter and Rhodovulum (Pfennig and Trüper, 1974b; Imhoff et al., 1984b; Hiraishi and Ueda, 1994a). The former are freshwater bacteria and the latter true marine bacteria. Species of both genera have distinct 16S rDNA sequences (Hiraishi and Ueda, 1994a; Hiraishi and Ueda, 1995a; Hiraishi et al., 1996). Two new species, Rhodovulum iodosum and Rhodovulum robiginosum, have been described that use ferrous iron as photosynthetic electron donor (Straub et al., 1999). Rhodobaca borogenensis, a new isolate from an alkaline soda lake with low salt concentrations, has adapted in its salt response to this habitat (Milford et al., 2000).

Habitats Ecological niches of phototrophic α-Proteobacteria are those anoxic parts of waters and sediments that receive light of sufficient quantity and quality to allow phototrophic development. Representatives of the purple nonsulfur bacteria are widely distributed in nature and are found not only in all kinds of stagnant water bodies, including lakes, waste water ponds, coastal lagoons, and in other aquatic habitats, but also in sediments, moist soils, and paddy fields. They live in aquatic habitats with significant amounts of soluble organic matter and low oxygen tension, but

CHAPTER 3.1.1

rarely form colored blooms, like those of purple sulfur bacteria. However, often they are found accompanying the purple sulfur bacteria in stratified environments. They have been found not only in freshwater, marine and hypersaline environments, and most frequently in habitats of moderate temperatures, but also in thermal springs and in cold polar habitats. Most purple nonsulfur bacteria have been isolated from all kinds of freshwater habitats where they also are most abundant. The greatest variety of species and the largest numbers of cells have been found in mud and water of eutrophic ponds, ditches and lakes. In the flat shore area of eutrophic lakes, 103 to more than 108 cells/ml of purple nonsulfur bacteria have been found in mud and water samples (Kaiser, 1966; Biebl and Drews, 1969). In pelagic water, the numbers usually are much lower (Biebl, 1973; Swoager and Lindstrom, 1971). Quite a number of purple nonsulfur bacteria occur in marine and hypersaline environments. They are usually found in water bodies and sediments of intertidal flats, salt marshes, and polluted harbor basins, but not in the open sea. While freshwater isolates have a very low tolerance to sulfide, the sulfide tolerance of most marine species is much higher, and they even use sulfide and thiosulfate as photosynthetic electron donors. This is certainly an adaptation to this environment, which characteristically has a high activity of sulfate reduction and where consequently anoxic conditions are coincident with the presence of hydrogen sulfide. Although some isolates (strains of Rhodopseudomonas palustris and Rhodomicrobium vannielii) from marine habitats are not typical marine bacteria and do not require salt for optimum growth, most of the purple nonsulfur bacteria found in marine habitats are typical marine bacteria and are not found in freshwater habitats (Imhoff, 1988a). The marine forms include species of the genera Rhodovulum and Rhodobium, as for example Rhodovulum sulfidophilum (Hansen and Veldkamp, 1973), Rhodovulum adriaticum (Neutzling et al., 1984), Rhodovulum euryhalinum (Kompantseva, 1985), Rhodobium marinum (Imhoff, 1983a) and Rhodobium orientis (Hiraishi et al., 1995b). Rhodospira trueperi and Roseospirillum parvum are from a marine salt marsh and also represent typical marine bacteria. In addition, Roseospira mediosalina was isolated from a hot spring with low salt concentrations (2% salts) and is growing optimally between 5 and 7% NaCl (Kompantseva and Gorlenko, 1984). Other halophilic species are well adapted to hypersaline environments. Rhodothalassium salexigens and Rhodovibrio salinarum are common to evaporated seawater pools and marine salt-

The Phototrophic Alpha-Proteobacteria

45

erns and sometimes form colored layers in salt deposits or sediments (Drews, 1981; Nissen and Dundas, 1984; Rodriguez-Valera et al., 1985), whereas Rhodovibrio sodomensis is from Dead Sea sediments (Mack et al., 1993). Some purple nonsulfur bacteria occur in acidic, boggy waters and soils. Most frequently, Rhodoblastus acidophilus, which grows optimally at pH 5.5–5.8, is found in such environments, often accompanied by Rhodopseudomonas palustris (Pfennig, 1969). Rhodopseudomonas palustris is very common and was isolated from all kinds of aquatic habitats (lakes, sewage and brackish waters), even from wet decaying leaves and from soils (Biebl and Pfennig, 1981; Imhoff and Trüper, 1992b). A preference for low pH values (pH 4.8-5.0) also is found in Rhodopila globiformis (Pfennig, 1974a). Purple nonsulfur bacteria are found regularly in all stages of conventional sewage plants. Their numbers may increase dramatically from raw sewage to the activated sludge stage (from 2,000 to 100,000 cells/ml; Siefert et al., 1978). When anaerobic sewage is incubated in the light, it often spontaneously becomes red-brown in color owing to the presence of phototrophic bacteria. This suggests that it should be possible to easily direct the processes in the activated sludge towards photoassimilation of organic substrates by phototrophic bacteria. The first sewage treatment plant based on this principle was established in Japan (Kobayashi et al., 1971; see below).

Isolation Two different strategies may be applied to isolate purple nonsulfur bacteria from their natural environment. Depending on the aims of the studies either the one or the other has clear preferences. According to classical enrichment techniques, phototrophic bacteria may be selectively enriched in suitable media under anoxic conditions and in the light; after visible growth, cells are separated in agar dilution series or on agar plates and isolated in pure culture. Alternatively pure cultures may be obtained by directly inoculating agar media from natural samples without prior enrichment. The first strategy is the method of choice whenever the isolation of bacteria with particular physiological features is attempted. With a proper environmental sample and the right medium, bacteria with the desired properties may be selected and isolated. The second strategy has to be used whenever the natural diversity is to be analyzed and information on the natural abundance and distribution of the species in a sample is required. While specifically adapted

46

J.F. Imhoff

media are essential for the success of the first strategy, in the second one nonselective media are used with substrates (such as acetate, succinate or malate) that enable the development of most purple nonsulfur bacteria (Biebl and Pfennig, 1981; Imhoff and Trüper, 1992b). Even if the medium used can support only poor growth of a specific strain, after separation in or on agar and when the incubation time is adapted to the slow growth rates, it will grow out to a small colony and can be picked up for further transfers.

Selective Enrichment Selective enrichment techniques for phototrophic bacteria were first used by Sergei Winogradsky (Winogradsky, 1888), and his technique is known as the “Winogradsky column.” Variations of this column technique by other authors are discussed by Pfennig (1965) and Van Niel (1971). Van Niel (1944) elaborated the physiological basis for the enrichment of purple nonsulfur bacteria and was the first to develop a defined medium which could be used for their enrichment and cultivation. For selective enrichment of purple nonsulfur bacteria, media have been used with lowered sulfate concentration to avoid production of sulfide by sulfate-reducing bacteria (Van Niel, 1944; Van Niel, 1971). An anaerobic enrichment culture using a common organic substrate and placed in the light usually will give rise to the development of purple nonsulfur bacteria. In many cases, in particular with marine samples, small Chromatiaceae—owing to their organotrophic capacity— compete with purple nonsulfur bacteria even if the sulfate concentration in the enrichment media is strongly reduced. The choice of the carbon source is not critical for the success of an enrichment culture because fermentative processes usually result in the formation of acetate or other acids (propionate, butyrate and lactate), which are good substrates for the majority of the purple nonsulfur bacteria. Some species of the purple nonsulfur bacteria, however, such as Rhodovulum sulfidophilum (Hansen and Veldkamp, 1973), Rhodovulum adriacticum (Neutzling et al., 1984) and Blastochloris sulfoviridis (Keppen and Gorlenko, 1975), tolerate and/or even require sulfide as a reduced sulfur source and/or photosynthetic electron donor. Besides the mineral salts composition and the concentration of nutrients of the media, also the addition of vitamins, the pH, temperature, light intensity and light regime are of general importance for the selectivity of enrichment cultures. If samples from marine and hypersaline environments are investigated, the salinity and the min-

CHAPTER 3.1.1

eral composition of the medium are of special importance. A few examples of selective parameters for the enrichment of purple nonsulfur bacteria are the following: Selective carbon sources for Rhodopseudomonas palustris are benzoate and formate (Qadri and Hoare, 1968). Because this species is very common in nature, many enrichments for purple nonsulfur bacteria will end up with the development of this species even without particularly selective conditions. Enrichments under photoautotrophic conditions with hydrogen as electron donor favor growth of Rhodobacter capsulatus, which grows faster under these conditions than other phototrophic bacteria do (Klemme, 1968). Also this species often becomes predominant in unselective media. Higher fatty acids like pelargonate and caproate (not more than 0.04% at pH 7.5) provide selective growth conditions for Phaeospirillum fulvum and Phaeospirillum molischianum (Pfennig, 1967; Van Niel, 1944). For enrichment of Rhodospirillum photometricum, the anaerobic infusion method of Molisch (Giesberger, 1947) is still the best method (Biebl and Pfennig, 1981). Hay or other dried plant materials are suitable sources which may be used in large test tubes or cylinders under continuous illumination. Rhodospirillum photometricum also can be readily enriched from activated sludge incubated anaerobically in the light (1,000 lux, 30°C; Siefert et al., 1978). The use of amino acids as carbon substrates favors the development of the Rhodospirillum species, in particular of Rhodospirillum rubrum (Biebl and Pfennig, 1981). A succinate-mineral salts medium without growth factors and with an initial pH of 5.5 is highly selective for both Rhodoblastus acidophilus and Rhodomicrobium vannielii (Pfennig, 1969). If yeast extract or the required vitamins are present Rhodopseudomonas palustris may also develop under acidic conditions. Enrichments in methanol bicarbonate medium select for Rhodoblastus acidophilus (and Rubrivivax gelatinosus). Typical marine bacteria will not or only poorly grow in media without NaCl, though most freshwater species are inhibited by NaCl concentrations of sea water. Therefore, the addition of 3% NaCl is a selective factor for marine purple nonsulfur bacteria. Salt concentrations of more than 10% are highly selective for moderately halophilic species like Rhodothalassium salexigens, Rhodovibrio salinarum and Rhodovibrio sodomensis that will not grow in media for freshwater or marine phototrophic bacteria.

CHAPTER 3.1.1

Isolation Procedures Direct Isolation. Methods of direct isolation of the phototrophic bacteria from a natural sample use agar dilution series or inoculation of agar plates to separate the cells prior to incubation. For isolation with solid media, water samples are most appropriate. A sample of mud, sludge, or soil may be used as a homogeneous suspension in medium or filter-sterilized water from the habitat. Samples containing less than 10 cells/ml need to be concentrated by centrifugation (agar dilution series) or filtration (agar plates). All methods for direct isolation are suitable for the determination of living cell counts, when known amounts of the sample are used in appropriate dilutions. Preparation of Agar Dilution Series. Agar dilution series are prepared either with enrichment cultures or with promising natural samples by direct inoculation with an environmental sample. Selective media are not required and nonselective ones are preferred for a direct isolation without prior enrichment procedure. In a modification of the method of Pfennig (Pfennig, 1965; Trüper, 1970), purified agar is dissolved (1.8%) and distributed in amounts of 3 ml into cotton-plugged test tubes. The agar is sterilized by autoclaving. The liquid agar is kept at 50°C in a water bath until use. A suitable medium is placed in the same water bath, and 6 ml of the prewarmed medium is added to each test tube. Medium and agar are mixed thoroughly by turning the tubes upside down and back and kept at 50°C. Eight tubes are sufficient for each dilution series. The first tube is inoculated with a natural sample or enrichment culture and mixed carefully; approximately 0.5 to 1.0 ml is transferred to a second tube, mixed carefully, and the procedure continued up to the last tube. The tubes are immediately placed into a cold water bath. After the agar has hardened, they are sealed with a paraffin mixture (3 parts paraffin oil and 1 part paraffin) and kept in the dark for several hours before they are incubated in the light (ca. 500–2,000 lux). After cells have grown to visible colonies, the paraffin layer is removed by melting, and well separated colonies are picked with a Pasteur pipette (the tip drawn out to a thin capillary) and transferred to a second dilution series. In general, three to four such dilution series are necessary to obtain pure cultures. When pure cultures have been obtained, single colonies are inoculated into liquid medium. Cultivation on Agar Plates. Purple nonsulfur bacteria have quite often been successfully isolated on agar plates. When high numbers of phototrophic bacteria are present in the sample, streaking by conventional methods is appropri-

The Phototrophic Alpha-Proteobacteria

47

ate. Samples containing low numbers of phototrophic bacteria can be easily concentrated on membrane filters (e.g., cellulose acetate or cellulose nitrate with 0.4 µm pore size; Biebl, 1973; Biebl and Drews, 1969; Swoager and Lindstrom, 1971; Westmacott and Primrose, 1975). The filter assembly is used after autoclaving, and the filters are transferred onto the agar surface with sterile tweezers. Samples with high numbers of purple nonsulfur bacteria (more than 100 cells/ml) may be diluted prior to filtration or distributed on the agar with a Drigalsky spatula (approximately 0.1 ml of a sample containing 100–300 cells/ml; Biebl and Drews, 1969). Incubation of the plates is recommended in an anaerobic jar in which the air is replaced by an oxygen-free mixture of nitrogen with 5% carbon dioxide and 3% hydrogen. Remaining traces of oxygen are removed by reaction with hydrogen over a palladium catalyst. Alternatively the GasPak system (Becton, Dickinson and Co.) or comparable systems may be used. A more detailed description of these methods is given by Biebl and Pfennig (1981). Normally, an incubation time of five or more days is required before intensely colored colonies of purple nonsulfur bacteria become visible. Occasionally, unicellular green algae develop and form flat spots of grass-green color. Sometimes purple sulfur bacteria, in particular Thiocapsa roseopersicina and Allochromatium vinosum, also develop.

Media A large number of media have been used for the enrichment and cultivation of purple nonsulfur bacteria (Biebl and Pfennig, 1981; Drews, 1965; Haskins and Kihara, 1967; Imhoff, 1982b; Imhoff and Trüper, 1976; Kaiser, 1966; Mack et al., 1993; Pfennig, 1969; Pfennig, 1978; Pratt and Gorham, 1970; Swoager and Lindstrom, 1971; Van Niel, 1944). For the isolation and cultivation of purple nonsulfur bacteria from freshwater and marine sources, the AT medium (Imhoff and Trüper, 1976; Imhoff, 1982b; Imhoff, 1988c) with numerous slight modifications has been successfully used as a basic medium for more than 20 years in the author’s laboratory. It can be used for cultivation of the great majority of the purple nonsulfur bacteria. It is also well suited for enrichment cultures of freshwater and marine purple nonsulfur bacteria. For the purpose of selective enrichment cultures, sulfate is omitted, a sulfate-free trace element solution (SLA) is used and selective carbon and nitrogen substrates are added. With nonselective carbon sources such as acetate, pyruvate, malate, succinate or fumarate, this medium is well suited for the isolation and enumeration of purple nonsul-

48

J.F. Imhoff

CHAPTER 3.1.1

fur bacteria in agar dilution series or on agar plates. A vitamin solution (VA) is used that adds sufficient amounts of all vitamins required by known purple nonsulfur bacteria (Imhoff, 1988c; see below). AT Medium Dissolve in and dilute to 1 liter with distilled water: KH2PO4 MgCl2 · 6H2O CaCl2 · 2H2O Na2SO4 NH4Cl NaHCO3 NaCl Na-acetate (or other carbon source)

1.0 g 0.5 g 0.1 g 0.7 g 1.0 g 3.0 g 1.0 g 1.0 g

Add: 1 ml of trace element solution SLA (see below) 1 ml of vitamin solution VA (see below) 10 ml of 5% sodium ascorbate Adjust pH to 6.9.

The medium is filter-sterilized, collected in an autoclaved 2-liter Erlenmeyer flask with an outlet at the bottom and distributed from this vessel into sterile screw-cap bottles of desired volume (usually 50-ml and 1-liter bottles) under sterile conditions. These bottles are filled completely (not more than a pea-sized air bubble should be left) and can be stored for several months. To achieve anoxic conditions and to remove the oxygen from the medium, 0.05% sodium ascorbate is added. To avoid oxidation of the ascorbate prior to distribution into the bottles, it is added to the medium immediately before sterile filtration. Yeast extract stimulates growth of most of the known purple nonsulfur bacteria. It is used as a source of growth factors and may be added at a concentration of 0.05%. For Phaeospirillum and Rhodospirillum species addition of 0.01% Fe-citrate is growth stimulating. It may also be routinely added to the medium, except when sulfide is present. For the cultivation of Rhodoblastus acidophilus and Rhodomicrobium vannielli, the pH is adjusted to 5.5. For marine species, NaCl is added. A saline modification of the medium, the SAT medium, contains 3% NaCl. For iron-oxidizing purple nonsulfur bacteria, 10 mM ferrous iron is added to completely anoxic media buffered with 20 mM bicarbonate at pH 7.0 (Ehrenreich and Widdel, 1994). An anoxic stock solution of FeSO4 is autoclaved, maintained under nitrogen, and used to supplement the media. To increase the solubility of iron, chelators such as citrate or nitrilotriacetic acid may be added. Some purple nonsulfur bacteria can tolerate sulfide to various degrees and/or use it as a photosynthetic electron donor. For these species,

sodium sulfide is added at low concentrations of 0.4–1.0 mM (up to 2 mM for more tolerant strains). Some species even are dependent on the presence of reduced sulfur sources because they lack the ability to use sulfate as an assimilatory sulfur source. For these bacteria also low concentrations of sulfide may be used, or alternative sources of reduced sulfur such as thiosulfate, cysteine or methionine may be applied. Vitamin Solution VA (Modified after Imhoff and Trüper, 1977) Biotin Niacin Thiamine dichloride p-Aminobenzoic acid Pyridoxolium hydrochloride Ca-panthothenate Vitamin B12

10 mg 35 mg 30 mg 20 mg 10 mg 10 mg 5 mg

This vitamin solution meets the vitamin requirements of all purple nonsulfur bacteria. More specifically, for the culture of individual strains or species, the vitamins may be added individually or in desired combinations at the indicated concentration. Dissolve in 100 ml of distilled water, sterilize by filtration, keep refrigerated, and add 1 ml per liter of medium. Trace Element Solution SLA (Modified after Imhoff and Trüper, 1977) FeCl2 · 4H2O CoCl2 · 6H2O NiCl2 · 6H2O CuCl2 · 2H2O MnCl2 · 4H2O ZnCl2 H3BO3 Na2MoO4 · 2H2O

1,800 mg 250 mg 10 mg 10 mg 70 mg 100 mg 500 mg 30 mg

These salts are dissolved separately in a total of 900 ml of double-distilled water; the solutions are mixed, the pH is adjusted to about 2–3 with 1 N HCl, and the final volume is brought to 1 liter. Medium for Rhodopila globiformis (Modified after Pfennig, 1974a) Mannitol Gluconate KH2PO4 NaCl CaCl2 · 2H2O MgCl2 · 6H2O NH4Cl Na-thiosulfate

1.5 g 1.5 g 0.4 g 0.4 g 0.05 g 0.4 g 0.4 g 0.2 g

Dissolve the above in and dilute up to 1 liter with distilled water. Add separately: 1 ml of SLA (or equivalent trace element solution) 1 ml of VA 5 ml of 0.1% Fe-citrate solution Adjust pH to 4.9.

CHAPTER 3.1.1

The Phototrophic Alpha-Proteobacteria

Complex Medium (Modified after Nissen and Dundas, 1984) MgCl2 · 6H2O KH2PO4 NaCl (depending on desired salinity) Yeast extract (Difco) Proteose peptone (Difco) Na-malate Trace element solution SLA

3.5 g 0.3 g 100 g 1.5 g 1.5 g 10 mM 1 ml

Dissolve the above in and dilute up to 1 liter of distilled water. Then adjust pH to 7.0. All strains of halophilic purple nonsulfur bacteria including our own isolates grow well in this medium. This and the complex medium described by Drews (1981) are suitable for growth of Rhodothalassium salexigens and Rhodovibrio salinarum. Synthetic Medium for Halophilic (Purple Nonsulfur) Phototrophic α-Proteobacteria (Modified from Imhoff, 1988c) NaHCO3 KH2PO4 KCl CaCl2 · 2H2O MgCl2 · 6H2O Na2SO4 NaCl (depending on desired salinity) Proline Acetate Pyruvate K3 citrate Glycine betaine Na-glutamate Vitamin solution VA (see above) Trace element solution SLA (see above)

3.9 g 0.5 g 1.0 g 0.05 g 3.5 g 1.0 g 40–150 g 5.0 mM 2.0 mM 2.0 mM 10.0 mM 10.0 mM 5.0 mM 1 ml 1 ml

Dissolve the above in and dilute up to 1 liter of distilled water. Adjust pH to 7.0. Sterilize by filtration. For special biochemical and physiological studies, synthetic media (such as this one) are required. The NaCl concentration is varied according to the desired salinity.

Identification Identification of new isolates can be obtained only by detailed studies of physiological and morphological properties together with chemotaxonomic information and genetic sequence data. To distinguish closely related strains and species of the same genus, often DNA-DNA hybridization studies are required. Because of the great diversity of the phototrophic α-Proteobacteria, a tentative assignment to one of the known genera often is much easier. Owing to the different physicochemical requirements of the species (e.g., salinity, pH and possibly temperature) and their physiological potential, the choice of the medium and the cul-

49

ture conditions for enrichment and isolation restrict the number of possible species that will be able to develop. Important additional information can be obtained from the color, size and consistency of the colonies on agar plates, the color of cell suspensions, and microscopic examinations. The shape of the cells, cell width and length, motility, mode of division, formation of cell aggregates, and presence of slime capsules are relevant properties that can be recognized in the light microscope. The cell morphology of a few representative species using light microscopy is shown in Fig. 4. Ultrathin sections under the electron microscope reveal the fine structure of the cells, in particular the type of the internal membrane system. In many species, the color of cell suspensions is indicative of the major type of carotenoids present (Schmidt, 1978). Spirilloxanthin as the major component gives a pink or red color, increasing amounts of additional rhodopin turn the color to brown-red, rhodopin without significant amounts of spirilloxanthin results in brown, okenone results in purple-red, and rhodopinal results in purple-violet. Carotenoids of the spheroidene series give colors from yellowish brown to brownish red (depending on the content of oxygenated derivatives formed in the presence of oxygen) and greenish to beige-brown under strongly reducing conditions. Carotenoidless mutants of Rhodospirillum rubrum and Rhodobium marinum are blue in color. In addition to the color of cell suspensions, absorption spectra yield preliminary information on the predominant bacteriochlorophylls and on the kind of bacteriochlorophyll-protein complexes. The carotenoids absorb at 480–550 nm. Absorption bands of bacteriochlorophyll a in vivo are at 380, 590–600, and 800–900 nm. Owing to the formation of different light-harvesting complexes, absorption spectra show a remarkable variation in the long wavelength range from 800–900 nm. Absorption spectra of whole cells are measured with cell suspensions washed twice in medium or appropriate salt solutions and then suspended in 60% sucrose solution (Biebl and Drews, 1969). Better results are often achieved by using isolated internal membranes suspended in buffer. For this purpose, it is sufficient to break the cells by ultrasonication or with a French press and to separate whole cells and large cell fragments from the internal membranes by centrifugation at 15,000 g. For the detailed description of a new bacterium, careful physiological studies are required, including the utilization of substrates, relations to oxygen, the ability to grow in darkness, respiratory and fermentative growth, vitamin requirement, and ranges and optima of salt concentration, pH and temperature.

50

J.F. Imhoff

CHAPTER 3.1.1 a

c

In addition to the phenotypic characterization, information on the genetic relatedness of a new isolate has to be obtained. To achieve a phylogenetic classification, 16S rDNA sequences are used and when the distinction of closely related strains and species is required (DeBont et al., 1981), DNA-DNA hybridization studies are performed. For the description of a new species, also the determination of the G+C content of the DNA is recommended. Also chemotaxonomic properties have been found to be quite helpful to identify and classify new isolates of phototrophic bacteria (Hiraishi et al., 1984; Imhoff, 1984a; Imhoff, 1988b; Imhoff et al., 1982c; Imhoff et al., 1984b; 1998; Imhoff and Bias-Imhoff, 1995b; Thiemann and Imhoff, 1996; Imhoff and Süling, 1996; Pfennig et al., 1997). In particular the ring structure and the isoprenoid chain length of respiratory quinones and the fatty acid composition of the cell membranes are quite useful in identification (Hiraishi et al., 1984; Imhoff, 1984a). The structure and composition of polar lipids and lipopolysaccharides also are diagnostic properties of high value (Imhoff and Bias-Imhoff, 1995b; Weckesser et al., 1995). The need for sophisticated analytical methods, however, makes these properties less accessible for diagnosis. Characteristic properties of presently known species of the phototrophic purple α-Proteobacteria that are of diagnostic value are shown in Tables 1–3. An arbitrary selection of outstanding properties of a few selected species follows:

Fig. 4. Morphology of Rhodospirillum rubrum (a); Rhodobacter sphaeroides (b); Rhodopseudomonas acidophilia (c); Rhodomicrobium vannielii (d), left: Vegetative cells with polyhedral exospores; phrase contrast micrographs. (Bar = 10 µm.)

b

d

Cultures of Blastochloris viridis and Blastochloris sulfoviridis are olive-green in color and contain bacteriochlorophyll β with an absorption maximum at 1020–1030 nm. Cultures of Rhodospira trueperi are peach colored and reveal unusually low long-wavelength absorption maxima of bacteriochlorophyll b at 986 nm. The in vivo absorption spectra of Rhodospirillum rubrum and Rhodobium marinum are characterized by an unusually low absorption maximum at approximately 803 nm and a single dominant peak without a shoulder (should be checked in derivative spectra) at 870–885 nm. This is taken as indication of the lack of peripheral light-harvesting complexes. Additional species with such spectra are Rhodovibrio sodomensis, Rhodobaca bogoriensis and Rhodocista centenaria. For Rhodomicrobium vannielii, the peritrichous flagellation of swarmer cells, the formation of cell aggregates connected by thin filamentous tubes and the formation of exospore-like cysts of moderate heat resistance are typical. Swarming motility on agar surfaces is characteristic for Rhodocista centenaria, which also forms desiccation- and heat-resistant cysts (Favinger et al., 1989). A definitive growth requirement for salt is found in the marine species of Rhodovulum and Rhodobium and also in Rhodospira trueperi, Roseospira mediosalina and Roseospirillum parvum.

+

+

Red

Motility

Color

Q-10, RQ-10

Major quinones

Q-8, RQ-8

Large

64.8–65.8

Fresh water

6.5–7.5

25–30

None

Q-9, MK-9

Small

64.3–65.3

Fresh water

7.3

25–30

None

−

−

paba

a

Brown

Q-9, MK-9

Small

60.5–64.8

Fresh water

7.3

30

None

−

−

aa

a

Brown

+

Stacks

Stacks +

0.7–1.0

0.5–0.7

Phs molischianum

Phaeospirillum Phs. fulvum

Q-9

o

69.9

Fresh water warm springs

6.8

40–45

None

o

+

b, B12

a

Pink

+

Lamellae

1.0–2.0

Rhodocista Rci. centenaria

0.8–0.9

0.6–0.7

Q-10, MK-10

None

67.4

Saltern

7.5–8.0

42

4–6% (3–24)

−

+

7

o

None

66.2–66.6

Salt lakes

Q-7, MK-7

o

65.7

o

o

71.2

7.9 Marine sediments

7.0–7.5

30

1–2

+

+

o

a

Pink

+

Lamellae

0.4–0.6

Roseospirillum Reo. parvum

Marine sediments

25–30

2% (0.5–5)

10–14% (8–18) 35–40

+

−

b, t, pan

b

Beige

+

Vesicles

0.6–0.8

Rhodospira Rsa. trueperi

o

+

a c, n

a

Pink

+

Vesicles

c, n

Red

+

Vesicles

Rhv. sodomensis

Rhodovibrio Rhv. salinarum

o

o

66.6

Salty springs

7

30–35

4–7% (0.5–15)

+

(+)

t, paba, n

a

Pink

+

Vesicles

0.8–1.0

Roseospira Ros. mediosalina

Q-10, MK-10

o

64.0

Saltern

6.6–7.4

40

6–8% (5–20)

Q-9/10, MK-9/10 RQ-9/10

Small

66.3

Fresh water acidic springs

4.8–5.0

30–35

None

−

(+)

+ −

b, paba

a

Purple-red

+

Vesicles

1.6–1.8

Rhodopila Rpi. globiformis

glu

a

Red

+

Lamellae

0.6–0.7

Rhodothalsassium Rts. salexigens

Symbols: +, positive in most strains; −, negative in most strains; +/−, variable in different strains; o, not determined; (+), weak growth or microaerobic growth only. Abbreviations: b, biotin; t, thiamine; n, niacin; paba, p-aminobenzoic acid; pan, pantothenate; glu, glutathione; ICM, internal cell membrane; Q-10, ubiquinone 10; MK-10, menaquinones 10; and RQ-10, rhodoquinone 10. a Mol%.

Large

Fresh water

63.8–65.8

6.8–7.0

Optimal pH

Habitat

Cytochrome c size

30–35

Optimal temperature

DNA base ratioa

None

−

+

Oxidation of sulfide

Salt requirement

−

+

Aerobic growth

a

n

a

b

Bacteriochlorophyll

Vitamins required

Brown

Stacks

0.8–1.0

Vesicles

Cell diameter (µm)

1.1–1.5

Rsp. photometricum

Rhodospirillum

Rsp. rubrum

ICM

Characteristic

Genus Species

Table 1. Characteristic properties of the phototrophic Rhodospirillum and related phototrophic purple nonsulfur bacteria.

CHAPTER 3.1.1 The Phototrophic Alpha-Proteobacteria 51

− Sulfide, So None

+/− H2, TS, Sulfide Paba (biotin)

63.5

68.8

Small Q-10, MK-10, RQ-10 62.2–66.8 (Bd)

− + + − − − −

None

B12, niacin, paba − o o o − o o o o

− H2

+

+ Red to orange-red a None 5.5–6.0 25–30

Lamellae

1.0–1.3 Sessile −

Rhodoblastus Rbl. acidophilus

o −

+

a None 6.8–7.2 40

+ o

Lamellae

1 Sessile +

Rps. cryptolactis

− + − + − + − o Q-10, RQ-10 66.8 (HPLC)

Niacin

+ TS

+

a None 7.0–7.5 30

+ Pink

Lamellae

1.0 Sessile −

− + − + − + − o Q-10, RQ-10 69.6–69.7 (HPLC)

Thiamine, paba

+ TS

+

a None 7.0 30–35

+ Pink

Lamellae

0.8–1.0 Tube +

Rpl. elegans

Rhodoplanes Rpl. roseus

Lamellae

0.5–0.9 Sessile −

− +/− − − (+) − − Small Q-9, MK-9 66.3–71.4 (Bd)

Biotin, paba

− −

(+)

− − − − + + + o Q-8/10, MK-7/8 67.8–68.4 (CA)

61.8–63.8 (Bd)

+ Small Q-10, RQ-10

− − +/− − −

o H2, Sulfide

+

+ Orange-brown to red a None 6.0 30

1.0–1.2 Tube Large cell aggregates Lamellae

Rhodomicrobium Rmi. vannielii

Biotin, None paba pyridoxine

− TS, Sulfide

(+)

+ + Green to Olive-green olive-green b b None None 6.5–7.0 7.0 25–30 28–30

Lamellae

0.6–0.9 Tube +

Blc. sulfoviridis

Blastochloris Blc. viridis

− − − − + + − o Q-10, MK-10 65.2–65.7 (HPLC)

Biotin, paba

+ TS

+

+ Pink to red a 4–5% 7.0–7.5 30–35

Lamellae

0.7–0.9 Sessile −/+

− +/− + − + − (+) o Q-10, MK-10 62.4–64.1 (HPLC)

o

− Sulfide

(+)

+ Pink to red a 1–5% 6.9–7.1 25–30

Lamellae

0.7–0.9 Sessile −

Rbi. marinum

Rhodobium Rbi. orientis

J.F. Imhoff

Symbols: +, positive in most strains; –, negative in most strains; +/−, variable in different strains; o, not determined; and (+), weak growth or microaerobic growth only. Abbreviations: paba, p-aminobenzoic acid; (biotin), biotin is required by some strains; Q-10, ubiquinone 10; MK-10, menaquinones 10; RQ-10, rhodoquinone 10; Bd, buoyant density; HPLC, high-pressure liquid chromatography; and TS, thiosulfate.

64.8–66.3 (Bd)

+

+

Mol% G+C of the DNA

a o 6.0 25–35

− − + − + o + o o

+ Pink

+ Brown-red to red a None 6.9 30–37

+ +/− − − − + + Large Q-10

Lamellae

1.0–1.5 Sessile +

Rps. julia

Rhodopseudomonas

Lamellae

0.6–0.9 Tube +

Rps. palustris

Utilization of: Benzoate Citrate Formate Tartrate Glucose Thiosulfate Sulfide Cytochrome c2 size Major quinones

Bacteriochlorophyll Salt requirement Optimum pH Optimum temperature (°C) Aerobic dark growth Denitrification Photoautotrophic growth with Vitamins required

Internal membrane system Motility Color of cultures

Characteristic Cell diameter (µm) Type of budding Rosette formation

Genus Species

Table 2. Characteristic properties of Rhodopseudomonas and related phototrophic purple nonsulfur bacteria.

52 CHAPTER 3.1.1

+

+

Sulfate assimilated

69.5–70.2 (HPLC)

Mol% G+C of DNA

65.3 (Bd)

− + − + + − + − o

b, n, t, B12

+

65.5–66.8 (Bd)

68.4–69.9 (Bd)

− + + + + + + − o

b, t, n

t, (b, n) + +/− − +/− − − + − o

+

S

o

+

−a

Binary fission

Vesicle

+

2.0–2.5

Rba. sphaeroides

+

S

o

+

−a

Binary fission

Vesicle

+

0.5–1.2

Rba. capsulatus

64.4–67.5 (Tm)

− − − − − − − + o

b, paba, t

+

S /sulfate

o

−

−

Binary fission

Vesicle

−

0.6–0.8

Rba. veldkampii

58–59

− − o + o − o o o

b, t, [B12]

+

o

o

1–2

Binary fission

Vesicle

+

0.8–1.0

Rhodobaca Rhb. bogoriensis

67.3–67.7 (HPLC)

+ + (+) − − − − + o

b, paba, t

+

Sulfate

+

0.8–1.0

Binary fission

Vesicle

+

0.6–1.0

Rhv. strictum

66.3–66.6 (HPLC)

+ − − +/− + +/− + + −

b, n, paba, t

+

Sulfate

+

1–6

Binary fission

Vesicle

+

0.6–0.9

Rhv. sulfidophilum

64.9–66.7 (Tm)

+ − − − + + − + −

b, t

−

S /sulfate

o

−

2.5–7.5

Binary fission

Vesicle

−

0.5–0.8

Rhv. adriaticum

62.1–68.6 (Tm)

+/− − − +/− + +/− + + −

b, n, paba, t

−

S /sulfate

o

−

0.5–12

Binary fission

Vesicle

+

0.7–1.0

Rhv. euryhalinum

Rhodovulum

− − o + + − + + + 69 (HPLC)

66 (HPLC)

b, n, B12

+

−

2.5–5

Binary fission

Vesicle

−

0.5–0.8

Rhv. robiginosum

− − o + − − + + +

b, n

+

−

2.5–5

Binary fission

Vesicle

−

0.5–0.8

Rhv. iodosum

Symbols: +, positive in most strains; −, negative in most strains; +/−, variable in different strains; o, not determined; and (+), weak growth or microaerobic growth only. Abbreviations: b, biotin; t, thiamine; n, niacin; paba, p-aminobenzoic acid; (biotin), biotin is required by some strains; [B12], vitamin B12 growth stimulating but not absolutely required; Bd, buoyant density; Tm, thermal denaturation; and TS, thiosulfate. a Optimal growth in the absence of NaCl, but able to grow at 3% NaCl.

+ o − + + o o − o

Utilization of: Formate Citrate Tartrate Mannitol Glycerol Ethanol Hydrogen Thiosulfate Ferrous iron

b, n, t

+

Aerobic dark growth

Vitamins required

S

Oxidation products of sulfide

−

−

−a

NaCl required (%)

o

Budding

Lamellae

−

Vesicle

0.6–0.8

+

Rba. blasticus

0.6–1.0

Rba. azotoformans

Binary fission

Cell division

Internal membrane system

Motility

Cell diameter (µm)

Characteristic

Genus Species

Rhodobacter

Table 3. Characteristic properties of Rhodobacter and related phototrophic purple nonsulfur bacteria.

CHAPTER 3.1.1 The Phototrophic Alpha-Proteobacteria 53

54

J.F. Imhoff

The tolerance of salt concentrations of more than 10% and growth under these conditions are characteristic for the halophilic species Rhodothalassium salexigens, Rhodovibrio salinarum and Rhodovibrio sodomensis.

Preservation For short-term preservation, well-grown cultures may be kept in closed, air-tight screw-cap bottles at 6–10°C in a refrigerator (or cold room) or at room temperature for several months, even years. Maintenance transfer of liquid cultures should occur at intervals of 2–6 months. In particular Rhodopseudomonas palustris and some Rhodobacter species have been shown to maintain viable cells over very long time periods. Stock cultures of the brown-colored Rhodospirillum and Phaeospirillum species and of other oxygen-sensitive species should be grown in the presence of 0.05% sodium ascorbate and transfers should be made every 1–2 months. Also, stock cultures of Blastochloris sulfoviridis and Rhodopila globiformis should be transferred more frequently. Stock cultures are incubated anaerobically in screw-cap 50-ml bottles in the described media at 25–30°C and 2,000 lux and then taken from the light during late exponential growth phase. Stock cultures also may be maintained for prolonged times in agar stab cultures or in appropriate dilutions of agar dilution series, sealed with paraffin and kept in the dark. For long-term storage, preservation in liquid nitrogen is recommended. Well grown cultures are supplemented with 50% dimethyl sulfoxide to give a final concentration of 5%, thoroughly mixed, dispensed in plastic ampoules, sealed and frozen in liquid nitrogen. Preservation and storage in liquid nitrogen are possible with all strains tested with either dimethylsulfoxide (5%) or glycerol (10%) as a protecting agent. Several purple nonsulfur bacteria were succsessfully preserved by freeze-drying (Biebl and Malik, 1976). The best protecting agent was skim milk (20%) supplemented with 10% sucrose, but most strains also survived in 10% sucrose alone. The brown-colored Phaeospirillum and Rhodospirillum species could not be lyophilized successfully (Biebl and Malik, 1976).

Physiology A comprehensive treatment of the various aspects of the physiology of purple nonsulfur bacteria including structure, function and genetics of the photosynthetic apparatus is found in various chapters of The Photosynthetic Bacteria (Clayton and Sistrom, 1978) and Anoxygenic

CHAPTER 3.1.1

Phototrophic Bacteria (Blankenship et al., 1995). A short overview on physiology and photosynthesis is given by Drews and Imhoff (1991). In the following, the basic principles and a few specific examples of metabolic properties of the phototrophic α-Proteobacteria are presented.

Photosynthesis Purple nonsulfur bacteria are anoxygenic phototrophic bacteria, growing phototrophically under anoxic conditions in the light without producing oxygen. All species can grow photoorganoheterotrophically using organic substrates as photosynthetic electron donors and carbon sources. Many representatives also grow under photolithoautotrophic conditions with reduced sulfur compounds, with hydrogen or ferrous iron ions as electron donor and CO2 as sole carbon source. Anoxygenic photosynthesis depends on the presence of a complex membrane-bound photosynthetic apparatus, which includes reaction center and light harvesting (antenna) pigmentprotein complexes. The proteins of reaction center and antenna noncovalently bind bacteriochlorophyll, carotenoids and other cofactors in stoichiometric ratios. Most purple nonsulfur bacteria have two antenna complexes. The complexes of the reaction center are surrounded by core antenna (a B870 or B890 antenna complex with bacteriochlorophyll a and a B1020 complex with bacteriochlorophyll b) and additional peripheral antenna (B800-850 and B800-820 complexes with bacteriochlorophyll a). Species such as Rhodospirillum rubrum and Rhodobium marinum lack peripheral antenna complexes and can be recognized by their minor absorption maxima at 803 nm. Most purple nonsulfur bacteria have one type of peripheral antenna complex (B800-850). The synthesis of multiple forms of peripheral antenna polypeptides with varying proportions under different growth conditions (as found in Rhodoblastus acidophilus and Rhodopseudomonas palustris) and the formation of different peripheral antenna complexes result in a complex pattern of absorption maxima that is only resolved in derivative spectra (Brunisholz and Zuber, 1992; Zuber and Cogdell, 1995). The principal function of the photosynthetic apparatus is the light-mediated excitation of a bacteriochlorophyll molecule in the reaction center followed by charge separation and resulting in electron transfer through the membrane. Most purple nonsulfur bacteria have an internal membrane system in which the photosynthetic apparatus is integrated. These internal membranes form vesicles, tubules or lamellar structures and are interconnected to the cytoplasmic

CHAPTER 3.1.1

membrane. They can be isolated by cell rupture and fractionated centrifugation. Quite characteristically, the production of photosynthetic pigments, pigment-protein complexes and of the photosynthetic membrane structures is suppressed by oxygen. A notable exception is Rhodocista centenaria (Yildiz et al., 1991). (Note: We distinguish between the anoxygenic phototrophic purple nonsulfur bacteria [which use photosynthesis as primary energy source and are well adapted to the anoxic way of life] and the aerobic bacteriochlorophyll-containing bacteria [which primarily gain energy by aerobic respiration but are unable to grow by anoxygenic photosynthesis under anoxic conditions]. The latter are treated elsewhere.)

Respiration Chemoorganoheterotrophic growth in the presence of oxygen is common among purple nonsulfur bacteria and most of the known species are facultatively chemotrophic. While some species are very sensitive to oxygen, others grow equally well under oxic conditions in the dark at the full oxygen tension of air. Also chemolithoautotrophic growth with hydrogen or reduced sulfur compounds as electron donors and oxygen as electron acceptor has been demonstrated (Madigan and Gest, 1979; Siefert and Pfennig, 1979). Under anoxic dark conditions, growth of several species is also supported by respiratory electron transport in the presence of nitrate, nitrite, and nitrous oxide. Denitrification is induced by nitrate, either in the dark or in the light, but is suppressed by oxygen. A single strain of Rhodobacter sphaeroides, described as a subspecies, Rhodobacter sphaeroides f. sp. denitrificans (Satoh et al., 1976), but later recognized as a regular strain of Rhodobacter sphaeroides (De Bont et al., 1981; Imhoff, 1989a), was the first phototrophic purple bacterium known to use nitrate as an electron acceptor under anoxic dark conditions. Later, nitrate reduction was found in additional strains of Rhodobacter sphaeroides (Michalski and Nicholas, 1988), in strains of Rhodopseudomonas palustris (Klemme et al., 1980), and in Rhodobacter capsulatus (McEwan et al., 1984). Some of these strains could not grow with nitrate as sole nitrogen source, but dinitrogen produced during denitrification might be assimilated under these growth conditions (Dunstan et al., 1982). Also, Rhodobacter azotoformans, Rhodobium orientis and Rhodoplanes roseus and Rhodoplanes elegans are able to denitrify (Hiraishi and Ueda, 1994b; Hiraishi et al., 1995b; Hiraishi et al., 1996). Anaerobic growth on sugars with dimethylsulfoxide (DMSO) or trimethylamine-N-oxide (TMAO) as an additional oxidant has been

The Phototrophic Alpha-Proteobacteria

55

observed first in Rhodobacter capsulatus (Yen and Marrs, 1977) and later also in other species. Energy generation during growth with fructose and TMAO was proposed to be due to anaerobic respiration (Schultz and Weaver, 1982), and the generation of a membrane potential under these conditions has been demonstrated (McEwan et al., 1983). Later it was questioned that a true electron transport chain is involved in DMSO and TMAO reduction. The physiological role of these external oxidants was discussed in detail (Ferguson et al., 1987; Zannoni, 1995).

Fermentation In the absence of external electron acceptors, a number of purple nonsulfur bacteria can use fermentative processes for energy generation (Uffen, 1978). During fermentative growth, substrates or storage products are oxidized incompletely, and reduced organic compounds as well as CO2 and H2 are produced. Rhodospirillum rubrum forms succinate, acetate, propionate, formate, CO2 and H2 during fermentation of fructose (Schön and Biedermann, 1973), whereas acetate, formate and equimolar amounts of CO2 and H2 are produced from pyruvate (Uffen, 1973). Rhodobacter capsulatus produces succinate, lactate, acetate, CO2 and H2 under identical conditions (Schultz and Weaver, 1982).

Carbon Metabolism Organic carbon sources have quite different functions under phototrophic, respiratory and fermentative metabolism. Under phototrophic growth conditions, they serve primarily as a source of cellular carbon, but in addition may function as an electron source for photosynthetic electron transport; in the presence of inorganic electron donors, they may be exclusively photoassimilated. During respiratory growth, the major part of the carbon sources is completely oxidized and only a minor fraction is assimilated into cell substance. Enzymatic reactions of the citric acid cycle involved in substrate oxidation would be expected at elevated levels under these conditions and indeed have been found to be increased during respiratory growth of Rhodobacter capsulatus (Beatty and Gest, 1981). During fermentation, substrates or storage products are oxidized incompletely on a large scale, and reduced organic compounds and hydrogen are excreted to achieve a redox balance of the cells. Most of the purple nonsulfur bacteria can use a variety of different organic carbon sources. Intermediates of the tricarboxylic acid cycle in addition to acetate and pyruvate are generally used. A number of purple nonsulfur bacteria use straight-chain saturated fatty acids with 5–18 car-

56

J.F. Imhoff

bon atoms (Janssen and Harfoot, 1987). Also organic acids, amino acids, alcohols and carbohydrates support growth of many of these bacteria. Carbon substrates such as citrate and aromatic compounds are used by a few species only. Citrate for instance is used by Rubrivivax gelatinosus (a β-Proteobacterium), Rhodobacter sphaeroides, Rhodobacter blasticus, Rhodovulum strictum, Rhodothalassium salexigens, Rhodoblastus acidophilus, Rhodoplanes roseus, Rhodoplanes elegans and by some strains of Blastochloris viridis, Rhodobium marinum and Rhodopseudomonas palustris. Some of these bacteria grow weakly with citrate. A small number of species (Rhodopseudomonas palustris, Rubrivivax gelatinosus, Phaeospirillum fulvum, Rhodocyclus purpureus, Rhodoblastus acidophilus and Rhodomicrobium vannielii) can grow at the expense of aromatic organic compounds, such as benzoate, 3hydroxybenzoate, 4-hydroxybenzoate, 1,3,5trihydroxybenzene and other hydroxylated, methoxylated aromatic acids and aldehydes (Dutton and Evans, 1978). The ability to use aromatic compounds is best developed in Rhodopseudomonas palustris. This species also metabolizes phenolic acids, phenylalkane carboxylates, 4-hydroxy-cinnamate and related compounds (Harwood and Gibson, 1988; Gibson and Harwood, 1995). Most of these compounds support both phototrophic (anoxic conditions in the light) and chemotrophic (oxic conditions in the dark) growth; some are degraded only under the one or the other condition. During phototrophic growth, a reductive pathway for cleavage of the aromatic ring structure is used (Dutton and Evans, 1969; Gibson and Harwood, 1995). Acetate is assimilated by almost all purple nonsulfur bacteria. The metabolic pathways of acetate assimilation, however, are quite different among the species. In many phototrophic purple bacteria, the primary reaction of acetate metabolism is the ATP-dependent formation of acetyl CoA, which is the substrate for further reactions. The two key enzymes of the glyoxylate cycle, isocitrate lyase and malate synthase, are present in most of these bacteria and acetate is assimilated via this pathway. There is, however, some variance in the literature regarding the presence of isocitrate lyase, in particular in Rhodospirillum rubrum and Rhodobacter sphaeroides (Tabita, 1995). Certainly, alternative pathways of acetate assimilation are possible. For Rhodospirillum rubrum, the conversion of acetate to oxalacetate by two carboxylation reactions from acetate to pyruvate and further to oxalacetate has been postulated (Buchanan et al., 1967). In Rubrivivax gelatinosus (a β-Proteobacterium), photoassimilation of acetate is possible via the

CHAPTER 3.1.1

serine-hydroxypyruvate pathway (Albers and Gottschalk, 1976). For purple nonsulfur bacteria, CO2 is an important carbon source. Under autotrophic growth conditions with CO2 as sole carbon source, the Calvin cycle with ribulose bisphosphate carboxylase (RubisCO) as key enzyme is employed (Tabita, 1995). This enzyme is well studied and constitutes a major fraction of the cellular protein in bacteria that grow with CO2 as sole carbon source and use the Calvin cycle. Also, CO2 is required under heterotrophic growth conditions during assimilation of several reduced organic substrates. A number of carboxylating enzyme activities are responsible for this “heterotrophic CO2 fixation” (see Kondratieva, 1979; Tabita, 1995). For instance, assimilation of propionate is connected with a carboxylation to succinate. Also, long-chain fatty acids and other highly reduced substrates such as methanol require CO2 to elevate the oxidation-reduction level of these substrates to that of the cell material. One-carbon compounds, such as methanol and formate, also are used by strains of a limited number of species. Rhodomicrobium vannielii, Rhodobium marinum, Rhodoblastus acidophilus, Rhodospirillum rubrum, Rubrivivax gelatinosus, Rhodocyclus tenuis and several Rhodobacter and Rhodovulum species use formate as carbon source. Reasonable growth rates with methanol were found only in strains of Rhodoblastus acidophilus (Douthit and Pfennig, 1976). Apparently, the RubisCO pathway is involved in carbon assimilation of Rhodoblastus acidophilus also during growth on methanol and formate; both substrates are used as electron donors and are oxidized to CO2, which in turn is assimilated (Quale and Pfennig, 1975; Sahm et al., 1976). Rubrivivax gelatinosus is able to grow anaerobically in the dark with CO as sole source as carbon and energy. Under these conditions, CO is transformed into CO2 and H2, and RubisCO could be involved in assimilation of the latter (Uffen, 1983). In Rhodospirillum rubrum, CO is used under anoxic conditions during phototrophic growth and induces an oxygensensitive CO dehydrogenase (Bonam et al., 1989).

Sulfur Metabolism The role of reduced sulfur compounds as photosynthetic electron donors for purple nonsulfur bacteria was realized together with the recognition of their importance in the marine environment. Purple nonsulfur bacteria vary greatly in their sulfur metabolism. Most purple nonsulfur bacteria, in particular freshwater species, are inhibited by sulfide even

CHAPTER 3.1.1

at low concentrations. Some species, however, are quite tolerant to this toxic compound and use it as a photosynthetic electron donor (Hansen and Imhoff, 1985; Hansen and van Gemerden, 1972; Imhoff, 1982a; Imhoff, 1983a; Neutzling et al., 1984). The tolerance of Rhodovulum sulfidophilum is high and comparable to that of Allochromatium vinosum. Rhodomicrobium vannielii tolerates concentrations of 2–3 mM, whereas growth of Rhodobacter capsulatus is completely inhibited at 2 mM. At low concentrations of sulfide (0.4–0.8 mM), however, growth of this species is quite rapid. Growth of Rhodopseudomonas palustris, as of most of the purple nonsulfur bacteria, is inhibited at concentrations as low as 0.5 mM sulfide (Hansen and van Gemerden, 1972). In general, small amounts of yeast extract in the media increase the tolerance towards sulfide. In addition to sulfide, some purple nonsulfur bacteria can use thiosulfate as an electron donor. Extracellular elemental sulfur is the final oxidation product during sulfide oxidation of a number of purple nonsulfur bacteria, such as Rhodospirillum rubrum, Rhodobacter capsulatus, Rhodobacter sphaeroides (Hansen and Veldkamp, 1973) and Roseospira mediosalina (Kompantseva and Gorlenko, 1984). Elemental sulfur occurs outside the cells. Only with Rhodopseudomonas julia elemental sulfur was microscopically observed outside as well as inside the cells (Kompantseva, 1989). Oxidation of elemental sulfur is found in a limited number of species, as in Rhodobacter veldkampii, Rhodovulum euryhalinum, Rhodovulum adriaticum and Rhodopseudomonas julia. In these species, elemental sulfur is an intermediate product during oxidation to sulfate (Hansen, 1974; Hansen and Imhoff, 1985; Neutzling et al., 1984; Kompantseva, 1985; Kompantseva, 1989). Sulfate is formed from sulfide without formation of elemental sulfur in Rhodovulum sulfidophilum, Rhodovulum strictum, Rhodopseudomonas palustris and Blastochloris sulfoviridis (Hansen, 1974; Hiraishi and Ueda, 1995a; Neutzling et al., 1985). Tetrathionate is the only oxidation product of Rhodomicrobium vannielii grown in a chemostat, but in batch culture, sulfide reacts with the tetrathionate formed, so that thiosulfate is the major product (together with minor amounts of elemental sulfur) accumulated under these conditions; sulfate is not formed under either condition (Hansen, 1974). With a few notable exceptions, the purple nonsulfur bacteria can use sulfate as an assimilatory sulfur source. Apparently two different pathways occur in these bacteria, involving either adenosine 5′-phosphosulfate (APS) or 3′-phosphoadenosine 5′-phosphosulfate (PAPS; Imhoff, 1982b). In the presence of reduced sulfur compounds,

The Phototrophic Alpha-Proteobacteria

57

these are preferentially assimilated and the energetically expensive assimilation of sulfate is repressed (Imhoff et al., 1983b). Common alternative assimilatory sulfur sources include sulfite, thiosulfate, cysteine, glutathione, methionine and sulfide. Rhodovulum adriaticum, Rhodovulum iodosum, Rhodovulum robiginosum, Rhodovulum euryhalinum, Blastochloris sulfoviridis, Rhodopseudomonas julia, Rhodobacter veldkampii and Roseospirillum parvum require reduced sulfur sources and are unable to assimilate sulfate (Keppen and Gorlenko, 1975; Kompantseva, 1985; Kompantseva, 1989; Neutzling et al., 1984; Hansen and Imhoff, 1985; Pfennig, 1974a; Straub et al., 1999; Glaeser and Overmann, 1999). Growth of Rhodopila globiformis is inhibited by high concentrations of sulfate although at low concentration this can serve as an assimilatory sulfur source (Imhoff et al., 1981). Also Rhodobium marinum grows poorly with sulfate as an assimilatory sulfur source, but much better in the presence of low concentrations of reduced sulfur sources.

Nitrogen Metabolism Ammonia, dinitrogen, and several organic nitrogen compounds (e.g., glutamate, aspartate or yeast extract) are the most appropriate nitrogen sources of most purple nonsulfur bacteria. Nitrate is assimilated only by a few species (strains of Rhodospirillum rubrum, Rhodobacter sphaeroides, Rhodobacter capsulatus and Rhodopseudomonas palustris) and growth yields with nitrate are considerably lower than with other nitrogen sources (Göbel, 1978; Imhoff, 1982a). Nitrate assimilation is inducible by nitrate but repressed by ammonia and glutamate. Assimilation of ammonia is strongly regulated and two different major pathways occur. The primary pathway in purple nonsulfur bacteria is via glutamine synthetase and glutamate synthase, which is active if cellular concentrations of ammonia are low and if growth is occurring at low concentrations of ammonia or on dinitrogen and nitrate (Drews and Imhoff, 1991). The second pathway present in most purple nonsulfur bacteria is via glutamate dehydrogenase, which is active when ammonia concentrations are high. The ability to fix dinitrogen is a common property of most phototrophic purple bacteria (Madigan, 1995). Dinitrogen fixation occurs under phototrophic and chemotrophic growth conditions, and the activity and expression of nitrogenase underlie a complex regulatory cascade (Drews and Imhoff, 1991; Haselkorn, 1986; Ludden and Roberts, 1995; Madigan, 1995). It has been most intensively studied in Rhodospirillum rubrum, where it was first discovered by Kamen and Gest (1949), and in Rhodobacter capsulatus.

58

J.F. Imhoff

Nitrogenase activity is repressed and inactivated by oxygen as in other dinitrogen-fixing bacteria. Its synthesis is derepressed at low concentrations of ammonia, i.e., under conditions of nitrogen limitation.

Hydrogen Metabolism A great number of phototrophic purple bacteria can photoproduce hydrogen. With dinitrogen, glutamate or aspartate as nitrogen source, a number of carbon substrates (lactate, acetate, butyrate, malate and others) may be completely transformed to CO2 and H2, and these in turn may serve as substrates for photoautotrophic growth. After consumption of the organic substrate, cell suspensions of Rhodobacter capsulatus that produced, e.g., H2 from lactate, rapidly started to consume the produced H2 (Kelley et al., 1977). Similar observations were obtained with Rhodospirillum rubrum (Schick, 1971). Photoevolution of H2 is catalyzed by nitrogenase, which is inhibited by ammonia, high concentrations of yeast extract and amino acids degraded to produce ammonia. The reaction is not reversible, insensitive to CO (a common inhibitor of hydrogenases) and independent of the partial pressure of H2. Hydrogen evolution in purple nonsulfur bacteria also occurs during fermentative growth under anoxic dark conditions. This hydrogen evolution is catalyzed by a reversible hydrogenase or by formate hydrogenlyase and underlies similar regulatory rules as in other fermenting bacteria, i.e., it is strongly inhibited by CO (Drews and Imhoff, 1991; Sasikala et al., 1993). Hydrogen is not only produced, but also serves as an excellent photosynthetic electron donor for many purple nonsulfur bacteria and enables these bacteria to grow photolithoautotrophically. Hydrogen uptake is catalyzed by a reversible, membrane-bound hydrogenase, which is induced by hydrogen and independent of the nitrogen source. The membrane-bound hydrogenase is not inhibited by ammonia, but strongly inhibited by CO. During growth conditions of dinitrogen fixation, this uptake hydrogenase recycles the hydrogen produced by nitrogenase, and mutants lacking this hydrogenase demonstrate an increased hydrogen production during nitrogen fixation (Drews and Imhoff, 1991).

Applications Most prominent examples of the application of phototrophic purple nonsulfur bacteria are their use in sewage treatment processes and for production of biomass, biopolymers and molecular hydrogen. They may be used as a source for cell-

CHAPTER 3.1.1

free systems performing photosynthesis and ATP formation and for the production of vitamins and other organic molecules. Sewage contains a complex mixture of small organic molecules that are good substrates for purple nonsulfur bacteria. Phototrophic bacteria are regularly found in conventional sewage treatment plants (Holm and Vennes, 1971; Siefert et al., 1978). Facultative chemotrophic purple phototrophic bacteria compete best under such conditions. Because light-driven energy generation enables them to use compounds produced during anaerobic degradation processes, these bacteria are good candidates for application to the final stages of sewage treatment. A highly advanced system using phototrophic bacteria in the purification of municipal and industrial waste water is that developed by Kobayashi (Kobayashi et al., 1971; Kobayashi and Tschan, 1973; Kobayashi, 1977; Kobayashi and Kobayashi, 1995). This process uses the natural sequence of aerobic and anaerobic degradation, followed by culture of anoxygenic phototrophic bacteria and of green algae in separated reaction tanks (bioreactors). Phototrophic bacteria used in Kobayashi’s system include Rhodopseudomonas palustris, Rhodobacter capsulatus, Rhodobacter sphaeroides and Rubrivivax gelatinosus. The use of phototrophic bacteria for sewage treatment was recently summarized by Kobayashi and Kobayashi (1995). A number of different sources and processes have been used to produce bacterial biomass of phototrophic bacteria. Animal wastes (Ensign, 1977; Sasaki et al., 1990), soybean wastes (Sasaki et al., 1981), wheat bran (Shipman et al., 1975), municipal and industrial waste waters (Kobayashi, 1977), and clarified effluents of a biogas plant (Vrati, 1984) have been used. The produced biomass of phototrophic bacteria is a valuable source of animal feed; it is rich in vitamins and in essential and sulfur-containing amino acids (Vrati, 1984) and has been used in plankton production, in the culture of shrimp, and as food for fish and chicken (Kobayashi, 1977; Mitsui, 1979). Addition of phototrophic bacterial cells to the food increased the survival of fish as well as the production and quality of hens’ eggs (Kobayashi and Tchan, 1973). With similar success the cell biomass of phototrophic bacteria has been used as fertilizer in agriculture (Kobayashi and Tchan, 1973). It may be used also for the production of hydrogen (Vrati and Verma, 1983; Bolliger et al., 1985), of biotin (Fillipi and Vennes, 1971), and of 5-aminolevulinic acid (Sasaki et al., 1990) and for other purposes. Under nitrogen starvation, almost all phototrophic bacteria are able to produce molecular hydrogen. This process is mainly due to hydrogen evolution from nitrogenase. A large number

CHAPTER 3.1.1

of substrates has been used by different research groups and with different purple nonsulfur bacteria to produce hydrogen (Kumazawa and Mitsui, 1982; Sasikala et al., 1993). Several attempts also have been made to produce hydrogen with immobilized cells of phototrophic bacteria (Francou and Vignais, 1984; Planchard et al., 1989; Vincenzini et al., 1982; von Felten et al., 1985; Weetall et al., 1981). Cells of Rhodospirillum rubrum (e.g., immobilized in a packed column) produced hydrogen with lactate as electron donor for 3,000 hours, with an activity loss of 60% after this time (von Felten et al., 1985). During the first days, mean rates of hydrogen production were 18–24 µl H2 per mg dry weight and hour. Also cell-free systems were applied in hydrogen production (Mitsui, 1975). A recent comprehensive discussion of the technology of hydrogen production from phototrophic purple bacteria is given by Sasikala et al. (1993). Poly-β-hydroxy-butyrate has been known for long as a storage product of phototrophic purple bacteria. In fact, an array of similar substances collectively termed “poly-3-hydroxyalkanoates” (PHAs) is accumulated, of which poly-3-hydroxybutyrate (PHB) is the most common. The PHAs occur as inclusion bodies of the cells visible in the light microscope and can account for a major fraction of the cell dry weight under appropriate growth conditions. Under strong nitrogen starvation or other conditions that restrict protein synthesis, excess carbon is converted into PHAs. In both Rhodospirillum rubrum and Rhodobacter sphaeroides, large amounts of storage PHAs can be synthesized under controlled growth conditions. Rhodobacter sphaeroides can produce more than 60% of its mass as PHAs of which 98% was PHB and the remainder was poly-3-hydroxyvalerate (Brandl et al., 1991). Rhodospirillum rubrum can produce a number of copolymers including an interesting tetrapolymer consisting of C4, C5 and C6 repeating units (Brandl et al., 1989). A flexible polymerase which uses a variety of substrates yielding PHAs with different properties may be useful for the production of biodegradable thermoplastic polyesters of commercial value (Fuller, 1995). A copolymer of PHB and poly-3hydroxyvalerate is commercially produced using cells of Ralstonia eutropha. A recent summary of aspects of biopolymer production by phototrophic purple bacteria is given by Fuller (1995).

Acknowledgement. The calculations for and the preparation of phylogenetic trees by Dr. J. Süling are kindly acknowledged.

The Phototrophic Alpha-Proteobacteria

59

Literature Cited Akiba, R., R. Usami, and K. Horikoshi. 1983. Rhodopseudomonas rutila, a new species of nonsulfur purple photosynthetic bacteria. Int. J. Syst. Bacteriol. 33:551– 556. Albers, H., and G. Gottschalk. 1976. Acetate metabolism in Rhodopseudomonas gelatinosa and several other Rhodospirillaceae. Arch. Microbial. 111:45–49. Beatty, J. T., and H. Gest. 1981. Biosynthetic and bioenergetic functions of citric acid cycle reactions in Rhodopseudomonas capsulata. J. Bacteriol. 148:585– 593. Biebl, H., and G. Drews. 1969. Das in-vivo Spektrum als taxonomisches Merkmal bei Untersuchungen zur Verbreitung von Athiorhodaceae. Zentralbl. Bakteriol. Parasitenkde. Infektionskr. Hyg. Abt. II, Orig. 123:425– 452. Biebl, H. 1973. Die Verbreitung der schwefelfreien Purpurbakterien im Plussee und anderen Seen Ostholsteins (Ph.D. thesis). University of Freiburg. FRG. Biebl, H., and R. A. Malik. 1976. Long term preservation of phototrophic bacteria. In: G. A. Codd and W. D. P. Stewart (Eds.) Proceedings of the Second International Symposium on Photosynthetic Prokaryotes. Dundee, Scotland. 31–33. Biebl, H., and N. Pfennig. 1981. Isolation of members of Rhodospirillaceae. In: M. P. Starr, H. Stolp, H. G. Trüper, A. Balows and H. G. Schlegel (Eds.) The Prokaryotes. Springer-Verlag. Berlin, 267–273. Blankenship, R. E., M. T. Madigan, and C. E. Bauer. 1995. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. Bolliger, R., H. Zürrer, and R. Bachofen. 1985. Photoproduction of molecular hydrogen from waste of a sugar refinery by photosynthetic bacteria. Appl. Microbiol. Biotech. 23:147–151. Bonam, D., L. Lehman, G. P. Roberts, and P. W. Ludden. 1989. Regulation of carbon monoxide dehydrogenase and hydrogenase in Rhodospirillum rubrum: Effects of CO and oxygen on synthesis and activity. J. Bacteriol. 171:3102–3107. Brandl, H., E. J. Knee, R. C. Fuller, R. A. Gross, and R. W. Lenz. 1989. The ability of the phototrophic bacterium Rhodospirillum rubrum to produce various poly (β-hydroxyalkanoates): Potential sources for biodegradable polyesters. Int. J. Biol. Macromol. 11:49–56. Brandl, H., R. A. Gross, R. W. Lenz, R. Lloyd, and R. C. Fuller. 1991. The accumulation of poly (3-hydroxyalkanoates) in Rhodobacter sphaeroides. Arch. Microbiol. 155:337–340. Brunisholz, R., and H. Zuber. 1992. Structure, function and organization of antenna polypeptides and antenna complexes from the three families of Rhodospirillaceae. Eur. J. Photochem. Photobiol. B. Biol. 15:113–140. Buchanan, B. B., M. C. W. Evans, and D. I. Arnon. 1967. Ferredoxin-dependent carbon assimilation in Rhodospirillum rubrum. Arch. Microbiol. 59:32–40. Burgess, J. G., R. Kawaguchi, T. Sakaguchi, R. H. Thornhill, and T. Matsunaga. 1993. Evolutionary relationships among Magnetospirillum strains inferred from phylogenetic analysis of 16S rDNA sequences. J. Bacteriol. 175:6689–6694.

60

J.F. Imhoff

Clayton, R. K., and W. R. Sistrom. 1978. In: R. K. Clayton, and W. R. Sistrom (Eds.) The Photosynthetic Bacteria. Plenum Press. New York , NY. DeBont, J. A. M., A. Scholten, and T. A. Hansen. 1981. DNADNA hybridization of Rhodopseudomonas capsulata, Rhodopseudomonas sphaerodides, and Rhodopseudomonas sulfidophila strains. Arch. Microbiol. 128:271–274. Douthit, H. A., and N. Pfennig. 1976. Isolation and growth rates of methanol utilizing Rhodospirillaceae. Arch. Microbiol. 107:233–234. Drews, G. 1965. Die Isolierung schwefelfreier Purpurbakterien. Zentralbl. Bakteriol. Parasitenkd., Infektionskd. und Hygiene, Abt. 1, Suppl. 1 Abt. 1, Suppl. 1:170– 178. Drews, G. 1981. Rhodospirillum salexigens, spec. nov., an obligatory halophilic phototrophic bacterium. Arch. Microbiol. 130:325–327. Drews, G., and J. F. Imhoff. 1991. Phototrophic purple bacteria. In: J. M. Shively and L. L. Barton (Eds.) Variations in Autotrophic Life. Academic Press. London, UK. 51– 97. Dunstan, R. H., B. C. Kelley, and D. J. D. Nicholas. 1982. Fixation of dinitrogen derived from denitrification of nitrate in a photosynthetic bacterium, Rhodospeudomonas sphaeroides forma sp. denitrificans. J. Bacteriol. 150:100–104. Dutton, P. L., and W. C. Evans. 1969. The metabolism of aromatic compounds by Rhodopseudomonas palustris: A new reductive method of aromatic ring metabolism. Biochem. J. 113:525–536. Dutton, P. L., and W. C. Evans. 1978. Metabolism of aromatic compounds by Rhodospirillaceae. In: R. K. Clayton and W. R. Sistrom (Eds.) The Photosynthetic Bacteria. Plenum Press. New York, NY. 719–726. Ehrenreich, A., and F. Widdel. 1994. Anaerobic oxidation of ferrous iron by purple bacteria, a new type of phototrophic metabolism. Appl. Environ. Microbiol. 60:4517–4526. Ensign, J. C. 1977. Biomass production from animal waste by photosynthetic bacteria. In: H. G. Schlegel and J. Barnea (Eds.) Microbiol Energy Conversion. Pergamon Press. Oxford, 455–482. Fani, R., C. Bandi, M. Bazzicalupo, M. T. Ceccherini, S. Fancelli, E. Gallori, L. Gerace, A. Grifoni, N. Miclaus, and G. Damiani. 1995. Phylogeny of the genus Azospirillum based on 16S rDNA sequence. FEMS Microbiol. Lett. 129:195–200. Favinger, J., R. Stadtwald, and H. Gest. 1989. Rhodospirillum centenum, sp. nov. a thermotolerant cyst-forming anoxygenic photosynthetic bacterium. Ant. v. Leeuwenhoek 55:291–296. Ferguson, S. J., J. B. Jackson, and A. G. McEvan. 1987. Anaerobic respiration in Rhodospirillaceae: Characterisation of pathways and evaluation of roles in redox balancing during photosynthesis. FEMS Microbiol. Rev. 46:117– 143. Filippi, G. M., and J. W. Vennes. 1971. Biotin production and utilization in a sewage treatment lagoon. Appl. Microbiol. 22:49–54. Francou, N., and P. M. Vignais. 1984. Hydrogen production by Rhodopseudomonas capsulata cells entrapped in carrageenan beads. Lett. 6:639–644. Fuller, R. C. 1995. Polyesters and photosynthetic bacteria: From lipid cellular inclusions to microbial thermoplastics. In: R. E. Blankenship, M. T. Madigan, and C. E.

CHAPTER 3.1.1 Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 1245–1256. Gibson, J., and C. S. Harwood. 1995. Degradation of aromatic compounds by nonsulfur purple bacteria. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 991–1003. Giesberger, G. 1947. Some observations on the culture, physiology and morphology of some brown-red Rhodospirillum-species. Ant. v. Leeuwenhoek 13:135–148. Glaeser, J., and J. Overmann. 1999. Selective enrichment and characterisation of Roseospirillum parvum, gen. nov. and sp. nov., a new purple nonsulfur bacterium with unusual light absorption properties. Arch. Microbiol. 171:405–416. Göbel, F. 1978. Quantum efficiencies of growth. In: R. K. Clayton and W. R. Sistrom (Eds.) The Photosynthetic Bacteria. Plenum Press. New York, NY. 907–925. Hansen, T. A., and H. van Gemerden. 1972. Sulfide utilization by purple nonsulfur bacteria. Arch. Microbiol. 86:49–56. Hansen, T. A., and H. Veldkamp. 1973. Rhodopseudomonas sulfidophila nov. spec., a new species of the purple nonsulfur bacteria. Arch. Microbiol. 92:45–58. Hansen, T. A. 1974. Sulfide als voor Rhodospirillaceae (Ph.D. thesis). University of Groningen. Groningen, The Netherlands. Hansen, T. A., and J. F. Imhoff. 1985. Rhodobacter veldkampii, a new species of phototrophic purple nonsulfur bacteria. Int. J. Syst. Bacteriol. 35:115–116. Harwood, C. S., and J. Gibson. 1988. Anaerobic and aerobic metabolism of diverse aromatic compounds by the photosynthetic bacterium Rhodopseudomonas palustris. Appl. Environ. Microbiol. 54:712–717. Haselkorn, R. 1986. Organization of the genes for nitrogen fixation in photosynthetic bacteria and cyanobacteria. Ann. Rev. Microbiol. 40:525–547. Haskins, E. F., and T. Kihara. 1967. The use of spectrophotometry in ecological investigation of the facultatively anaerobic purple photosynthetic bacteria. Can. J. Microbiol. 13:1238–1293. Hiraishi, A., Y. Hoshino, and H. Kitamura. 1984. Isoprenoid quinone composition in the classification of Rhodospirillaceae. J. Gen. Appl. Microbiol. 30:197–210. Hiraishi, A., T. S. Santos, J. Sugiyama, and K. Komagata. 1992. Rhodopseudomonas rutila is a later subjective synonym of Rhodopseudomonas palustris. Int. J. Syst. Bacteriol. 42:186–188. Hiraishi, A., and Y. Ueda. 1994a. Intrageneric structure of the genus Rhodobacter: Transfer of Rhodobacter sulfidophilus and related marine species to the genus Rhodovulvum gen. nov. Int. J. Syst. Bacteriol. 44:15–23. Hiraishi, A., and Y. Ueda. 1994b. Rhodoplanes gen. nov., a new genus of phototrophic bacteria including Rhodopseudomonas rosea as Rhodoplanes roseus comb. nov. and Rhodoplanes elegans sp. nov. Int. J. Syst. Bacteriol. 44:665–673. Hiraishi, A., and Y. Ueda. 1995a. Isolation and characterization of Rhodovulum strictum sp. nov. and some other members of purple nonsulfur bacteria from colored blooms in tidal and seawater pools. Int. J. Syst. Bacteriol. 45:319–326. Hiraishi, A., K. Urata, and T. Satoh. 1995b. A new genus of marine budding phototrophic bacteria, Rhodobium gen. nov., which includes Rhodobium orientis sp. nov. and

CHAPTER 3.1.1 Rhodobium marinum comb. nov. Int. J. Syst. Bacteriol. 45:226–234. Hiraishi, A., K. Muramatsu, and Y. Ueda. 1996. Molecular genetic analyses of Rhodobacter azotoformans sp. nov. and related species of phototrophic bacteria. Syst. Appl. Microbiol. 19:168–177. Hiraishi, A. 1997. Transfer of the bacteriochlorophyll bcontaining phototrophic bacteria Rhodopseudomonas viridis and Rhodopseudomonas sulfoviridis to the genus Blastochloris gen. nov. Int. J. Syst. Bacteriol. 47:217–219. Holm, H. W., and J. W. Vennes. 1971. Occurrence of purple sulfur bacteria in a sewage treatment lagoon. Appl. Microbiol. 19:988–996. Imhoff, J. F., and H. G. Trüper. 1976. Marine sponges as habitats of anaerobic phototrophic bacteria. Microbial Ecol. 3:1–9. Imhoff, J. F., and H. G. Trüper. 1977. Ectothiorhodospira halochloris sp. nov., a new extremely halophilic phototrophic bacterium containing bacteriochlorophyll b. Arch. Microbiol. 114:115–121. Imhoff, J. F., J. Then, F. Hashwa, and H. G. Trüper. 1981. Sulfate assimilation in Rhodopseudomonas globiformis. Arch. Microbiol. 130:234–237. Imhoff, J. F. 1982a. Response of photosynthetic bacteria to mineral nutrients. In: A. Mitsui and C. C. Black (Eds.) CRC Handbook of Biosolar Resources, Volume 1: Basic Principles, Part 1. CRC Press. Boca Raton, FL. 135–146. Imhoff, J. F. 1982b. Occurrence and evolutionary significance of two sulfate assimilation pathways in the Rhodospirillaceae. Arch. Microbiol. 132:197–203. Imhoff, J. F., D. J. Kushner, S. C. Kushawa, and M. Kates. 1982c. Polar lipids in phototrophic bacteria of the Rhodospirillaceae and Chromatiaceae families. J. Bacteriol. 150:1192–1201. Imhoff, J. F. 1983a. Rhodopseudomonas marina sp. nov., a new marine phototrophic purple bacterium. Syst. Appl. Microbiol. 4:512–521. Imhoff, J. F., M. Kramer, and H. G. Trüper. 1983b. Sulfate assimilation in Rhodopseudomonas sulfidophila. Arch. Microbiol. 136:96–101. Imhoff, J. F. 1984a. Quinones of phototrophic purple bacteria. FEMS Microbiol. Lett. 25:85–89. Imhoff, J. F., H. G. Trüper, and N. Pfennig. 1984b. Rearrangements of the species and genera of the phototrophic “purple nonsulfur bacteria.” Int. J. Syst. Bacteriol. 34:340–343. Imhoff, J. F. 1988a. Halophilic phototrophic bacteria. In: F. Rodriguez-Valera (Ed.) Halophilic Bacteria. CRC Press. Boca Raton, FL. 85–108. Imhoff, J. F. 1988b. Lipids, fatty acids and quinones in taxonomy and phylogeny of anoxygenic phototrophic bacteria. In: J. M. Olson, J. G. Ormerod, J. Amesz, E. Stackebrandt and H. G. Trüper (Eds.) Green Photosynthetic Bacteria. Plenum Press. New York, NY. 223–232. Imhoff, J. F. 1988c. Anoxygenic phototrophic bacteria. In: B. Austin (Ed.) Methods in Aquatic Bacteriology. Wiley and Sons. Chichester, UK. Imhoff, J. F. 1989a. Genus Rhodobacter. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 3:1668–1672. Imhoff, J. F., and H. G. Trüper. 1989b. The purple nonsulfur bacteria. In: J. T. Staley, M. P. Bryant, N. Pfennig and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 3:1658– 1661.

The Phototrophic Alpha-Proteobacteria

61

Imhoff, J. F. 1992a. Taxonomy, phylogeny and general ecology of anoxygenic phototrophic bacteria. In: N. G. Carr and N. H. Mann (Eds.) Biotechnology Handbook: Photosynthetic Prokaryotes. Plenum Press. London, New York, NY. 53–92. Imhoff, J. F., and H. G. Trüper. 1992b. The genus Rhodospirillum and related genera. In: A. Balows, H. G. Trüper, M. Dworkin, W. Harder and K. H. Schleifer (Eds.) The Prokaryotes (2nd ed.). Springer-Verlag. New York, NY. 2141–2155. Imhoff, J. F. 1995a. Taxonomy and physiology of phototrophic purple bacteria and green sulfur bacteria. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 1– 15. Imhoff, J. F., and U. Bias-Imhoff. 1995b. Lipids, quinones and fatty acids of anoxygenic phototrophic bacteria. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 179–205. Imhoff, J. F., and J. Süling. 1996. The phylogenetic relationship among Ectothiorhodospiraceae: A reevaluation of their taxonomy on the basis of rDNA analyses. Arch. Microbiol. 165:106–113. Imhoff, J. F., R. Petri, and J. Süling. 1998. Reclassification of species of the spiral-shaped phototrophic purple nonsulfur bacteria of the alpha-Proteobacteria: Description of the new genera Phaeospirillum gen. nov., Rhodovibrio gen. nov., Rhodothalassium gen. nov. and Roseospira gen. nov. as well as transfer of Rhodospirillum fulvum to Phaeospirillum fulvum comb. nov., of Rhodospirillum molischianum to Phaeospirillum molischianum comb. nov., of Rhodospirillum salinarum to Rhodovibrio salinarum comb. nov., of Rhodospirillum sodomense to Rhodovibrio sodomensis comb. nov., of Rhodospirillum salexigens to Rhodothalassium salexigens comb. nov., and of Rhodospirillum mediosalinum to Roseospira mediosalina comb. nov. Int. J. Syst. Bacteriol. 48:957– 964. Imhoff, J. F. 1999. A phylogenetically oriented taxonomy of anoxygenic phototrophic bacteria. In: G. A. Pescheck, W. Löffelhardt, and G. Schmetterer (Eds.) The Phototrophic Prokaryotes. Plenum Press. New York, NY. 763–774. Imhoff, J. F. 2000. The anoxygenic phototrophic purple bacteria. In: D. R. Boone and R. W. Castenholz (Eds.) Bergey’s Manual of Systematic Bacteriology, 2nd ed. Springer-Verlag. New York, NY. 1. Imhoff, J. F. 2001. Taxonomic note: Transfer of Rhodopseudomonas acidophila to the new genus Rhodoblastus as Rhodoblastus acidophilus comb. nov. Int. J. Syst. Evol. Microbiol. Janssen, P. H., and C. G. Harfoot. 1987. Phototrophic growth on n-fatty acids by members of the family Rhodospirillaceae. Syst. Appl. Microbiol. 9:9–11. Kaiser, P. 1966. Contribution a l’étude de l’écologie des bacteries photosynthetiques. Ann. Inst. Pasteur 111:733– 749. Kamen, M. D., and H. Gest. 1949. Evidence for a nitrogenase system in the photosynthetic bacterium Rhodospirillum rubrum. Science 109:560. Kawasaki, H., Y. Hoshino, H. Kuraishi, and K. Yamasato. 1992. Rhodocista centenaria gen. nov., sp. nov., a cystforming anoxygenic photosynthetic bacterium and its

62

J.F. Imhoff

phylogenetic position in the Proteobacteria alpha group. J. Gen. Appl. Microbiol. 38:541–551. Kawasaki, H., Y. Hoshino, and K. Yamasato. 1993a. Phylogenetic diversity of phototrophic purple non-sulfur bacteria in the Proteobacteria alpha-group. FEMS Microbiol. Lett. 112:61–66. Kawasaki, H., Y. Hoshino, A. Hirata, and K. Yamasato. 1993b. Is intracytoplasmic membrane structure a generic criterion? It does not coincide with phylogenetic interrelationships among phototrophic purple nonsulfur bacteria. Arch. Microbiol. 160:358–362. Kelley, B. C., C. M. Meyer, C. Gandy, and P. M. Vignais. 1977. Hydrogen recycling by Rhodopseudomonas capsulata. FEBS Lett. 81:281–285. Keppen, O. I., and V. M. Gorlenko. 1975. A new species of purple budding bacteria containing bacteriochlorophyll b. Microbiologiya 44:258–264. Klemme, J. H. 1968. Untersuchungen zur Photoautotrophie mit molekularem Wasserstoff bei neuisolierten schwefelfreien Purpurbakterien. Arch. Microbiol. 64:29–42. Kobayashi, M., M. Kobayashi, and H. Nakanishi. 1971. Construction of a purification plant for polluted water using photosynthetic bacteria. J. Ferment. Technol. 49:817– 825. Kobayashi, M., and Y. T. Tchan. 1973. Treatment of industrial waste solutions and production of useful byproducts using photosynthetic bacterial method. Water Res. 7:1219–1224. Kobayashi, M. 1977. Utilization and disposal of wastes by photosynthetic bacteria. In: H. G. Schlegel and J. Barnea (Eds.) Microbial Energy Conversion. Pergamon Press. Oxford, 443–453. Kobayashi, M., and M. Kobayashi. 1995. Waste remediation and treatment using anoxygenic phototrophic bacteria. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 1269–1282. Kompantseva, E. I., and V. M. Gorlenko. 1984. A new species of moderately halophilic purple bacterium Rhodospirillum mediosalinum sp. nov. (English translation). Mikrobiologiya 53:775–781. Kompantseva, E. I. 1985. Rhodobacter euryhalinus new species a new halophilic purple bacterial species. Mikrobiologiya 54:974–981. Kompantseva, E. I. 1989. A new species of budding purple bacteria: Rhodopseudomonas julia sp. nov. (English translation). Mikrobiologiya 58:254–259. Kondratieva, E. N. 1979. Interrelation between modes of carbon assimilation and energy production in phototrophic purple and green bacteria. In: J. R. Quale (Ed.) Microbial Biochemistry: International Review of Biochemistry. University Park Press. Baltimore, MD. 21:117–175. Kumazawa, S., and A. Mitsui. 1982. Hydrogen metabolism of photosynthetic bacteria and algae. In: A. Mitsui and C. C. Black (Eds.) CRC Handbook of Biosolar Resources, Volume 1: Basic Principles, Part 1. CRC Press. Boca Raton, FL. 299–316. Klemme, J. H., I. Chyla, and M. Preuss. 1980. Dissimilatory reduction by strains of the faculative phototrophic bacterium Rhodopseudomonas palustris. FEMS Microbiol. Lett. 9:137–140. Ludden, P. W., and G. P. Roberts. 1995. The biochemistry and genetics of nitrogen fixation by photosynthetic bacteria. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer

CHAPTER 3.1.1 (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 929– 947. Mack, E. E., L. Mandelco, C. R. Woese, and M. T. Madigan. 1993. Rhodospirillum sodomense, sp. nov. a Dead Sea Rhodospirillum species. Arch. Microbiol. 160:363– 371. Madigan, M. T., and H. Gest. 1979. Growth of the photosynthetic bacterium Rhodopseudomonas capsulata chemoautotrophically in the darkness with H2 as energy source. J. Bacteriol. 137:524–530. Madigan, M. T. 1995. Microbiology of nitrogen fixation by anoxygenic photosynthetic bacteria. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 915–928. McEwan, A. G., S. J. Ferguson, and J. B. Jackson. 1983. Electron flow to dimethylsulphoxide or trimethylamineN-oxide generates a membrane potential in Rhodopseudomonas capsulata. Arch. Microbiol. 136:300–305. McEwan, A. G., J. B. Jackson, and S. J. Ferguson. 1984. Rationalization of properties of nitrate reductases in Rhodopseudomonas capsulata. Arch. Microbiol. 137:344–349. Michalski, W. P., and D. J. D. Nicholas. 1988. Identification of two new denitrifying strains of Rhodobacter sphaeroides. FEMS Microbiol. Lett. 52:239–244. Migula, W. 1900. System der Bakterien. Gustav Fischer. Jena, Germany. 2. Milford, A. D., L. A. Aschenbach, D. O. Jung, and M. T. Madigan. 2000. Rhodobaca bogoriensis gen. nov. and sp. nov., an alkaliphilic purple nonsulfur bacterium from African Rift Valley soda lakes. Arch. Microbiol. 174:18– 27. Mitsui, A. 1975. The utilization of solar energy for hydrogen production by cell free system of photosynthetic organisms. In: T. N. Veziroglu (Ed.) Hydrogen Energy. Plenum Press. New York, NY. 309–316. Mitsui, A. 1979. Biosaline research. In: A. Hollaender, J. C. Aller, E. Epstein, A. San Pietro and O. Zaborsky (Eds.) The Use of Photosynthetic Marine Organisms in Food and Feed Production. Plenum Press. New York, NY. 177–215. Molisch, H. 1907. Die Purpurbakterien nach neuen Untersuchungen. G. Fischer. Jena, Germany. 1–95. Neutzling, O., J. F. Imhoff, and H. G. Trüper. 1984. Rhodopseudomomas adriatica sp. nov., a new species of the Rhodospirillaceae, dependent on reduced sulfur compounds. Arch. Microbiol. 137:256–261. Neutzling, O., C. Pfleiderer, and H. G. Trüper. 1985. Dissimilatory sulphur metabolism in phototrophic “nonsulphur” bacteria. J. Gen. Microbiol. 131:791–798. Nissen, H., and I. D. Dundas. 1984. Rhodospirillum salinarum sp. nov., a halophilic photosynthetic bacterium from a Portuguese saltern. Arch. Microbiol. 138:251– 256. Pfennig, N. 1965. Anreicherungskulturen für rote und grüne Schwefelbakterien. Zentralbl. Bakteriol. Parasitenkd. Infektionskrankh. Hyg. Abt. 1, Orig. Suppl. 1 179–189, 503–505. Pfennig, N. 1967. Photosynthetic bacteria. Ann. Rev. Microbiol. 21:285–324. Pfennig, N. 1969. Rhodospeudomonas acidophila, sp. n., a new species of the budding purple nonsulfur bacteria. J. Bacteriol. 99:597–602.

CHAPTER 3.1.1 Pfennig, N., and H. G. Trüper. 1971. Higher taxa of the phototrophic bacteria. Int. J. Syst. Bacteriol. 21:17–18. Pfennig, N. 1974a. Rhodopseudomonas globiformis, sp. n., a new species of the Rhodospirillaceae. Arch. Microbiol. 100:197–206. Pfennig, N., and H. G. Trüper. 1974b. The phototrophic bacteria. In: R. E. Buchanan and N. E. Gibbons (Eds.) Bergey’s Manual of Determinative Bacteriology, 8th ed. Williams and Wilkins. Baltimore, MD. 24–75. Pfennig, N. 1978. Rhodocyclus purpureus gen. nov. and sp. nov., a ring-shaped vitamin B12-requiring member of the family Rhodospirillaceae. Int. J. Syst. Bacteriol. 28:283– 288. Pfennig, N., H. Lünsdorf, J. Süling, and J. F. Imhoff. 1997. Rhodospira trueperi, gen. nov. and spec. nov., a new phototrophic Proteobacterium of the alpha-group. Arch. Microbiol. 168:39–45. Planchard, A., L. Mignot, T. Jouenne, and G.-A. Junter. 1989. Photoproduction of molecular hydrogen by Rhodospirillum rubrum immobilized in composite agar layer/ microporous membrane structures. Appl. Microbiol. Biotechnol. 31:49–54. Pratt, D. C., and E. Gorham. 1970. Occurrence of Athiorhodaceae in woodland, swamp, and pond soils. Ecology 51:346–349. Qadri, S. M. H., and D. S. Hoare. 1968. Formic hydrogenlyase and the photoassimilation of formate by a strain of Rhodopseudomonas palustris. J. Bacteriol. 95:2344– 2357. Quale, J. R., and N. Pfennig. 1975. Utilization of methanol by Rhodospirillaceae. Arch. Microbiol. 102:193– 198. Rodriguez-Valera, F., A. Ventosa, G. Juez, and J. F. Imhoff. 1985. Variation of environmental features and microbial populations with salt concentrations in a multipond saltern. Microbial Ecol. 11:107–115. Sahm, J., R. B. Cox, and J. R. Quale. 1976. Metabolism of methanol by Rhodopseudomonas acidophila. J. Gen. Microbiol. 94:313–322. Sasaki, K., N. Noparatnaraporn, M. Hayashi, Y. Nishizawa, and S. Nagai. 1981. Single-cell protein production by treatment of soybean wastes with Rhodopseudomonas gelatinosa. J. Ferm. Technol. 59:471–477. Sasaki, K., T. Tanaka, Y. Nishizawa, and M. Hayashi. 1990. Production of a herbicide, 5-aminolevulinic acid, by Rhodobacter sphaeroides using the effluent of swine waste from an anaerobic digestor. Appl. Microbiol. Biotechnol. 32:727–731. Sasikala, K., C. V. Ramana, P. R. Rao, and K. L. Kovacs. 1993. Anoxygenic phototrophic bacteria: Physiology and advances in hydrogen production technology. Adv. Appl. Microbiol. 38:211–295. Satoh, T., Y. Hoshino, and H. Kitamura. 1976. Rhodopseudomonas sphaeroides f sp. denitrificans, a denitrifying strain as a subspecies of Rhodopseudomonas sphaeroides. Arch. Microbiol. 108:265–269. Schick, H. J. 1971. Interrelationship of nitrogen fixation, hydrogen evolution and photoreduction in Rhodospirillum rubrum. Arch. Microbiol. 75:102–109. Schmidt, K. 1978. Biosynthesis of carotenoids. In: R. K. Clayton and W. R. Sistrom (Eds.) The Photosynthetic Bacteria. Plenum Press. New York, NY. 729–750. Schön, G., and M. Biedermann. 1973. Growth and adaptive hydrogen production of Rhodospirillum rubrum (F1) in anaerobic dark cultures. Biochim. Biophys. Acta 304:65– 75.

The Phototrophic Alpha-Proteobacteria

63

Schultz, J. E., and P. F. Weaver. 1982. Fermentation and anaerobic respiration by Rhodospirillum rubrum and Rhodopseudomonas capsulata. J. Bacteriol. 149:181– 190. Seewaldt, E., K.-H. Schleifer, E. Bock, and E. Stackebrandt. 1982. The close phylogenetic relationship of Nitrobacter and Rhodopseudomonas palustris. Arch. Microbiol. 131:287–290. Shipman, R. H., I. C. Kao, and L. T. Fan. 1975. Single-cell protein production by photosynthetic bacteria cultivation in agricultural by-products. Biotechnol. Bioeng. 17:1561–1570. Siefert, E., R. L. Irgens, and N. Pfennig. 1978. Phototrophic purple and green bacteria in a sewage treatment plant. Appl. Environ. Microbiol. 35:38–44. Siefert, E., and N. Pfennig. 1979. Chemoautotrophic growth of Rhodopseudomonas species with hydrogen and chemotrophic utilization of methanol and formate. Arch. Microbiol. 122:177–182. Sievers, M., W. Ludwig, and M. Teuber. 1994. Phylogenetic positioning of Acetobacter, Gluconobacter, Rhodopila and Acidiphilium species as a branch of acidophilic bacteria in the alpha-subclass of proteobacteria based on 16S ribosomal DNA sequences. Syst. Appl. Microbiol. 17:189–196. Stackebrandt, E., R. G. E. Murray, and H. G. Trüper. 1988. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “purple bacteria and their relatives.” Int. J. Syst. Bacteriol. 38:321–325. Stadtwald-Demchick, R., F. R. Turner, and H. Gest. 1990. Rhodopseudomonas cryptolactis, sp. nov., a new thermotolerant species of budding phototrophic purple bacteria. FEMS Microbiol. Lett. 71:117–122. Straub, K. L., F. A. Rainey, and F. Widdel. 1999. Rhodovulum iodosum sp. nov. and Rhodovulum robiginosum sp. nov., two new marine phototrophic ferrous-iron-oxidizing purple bacteria. Int. J. Syst. Bacteriol. 49:729–735. Swoager, W. C., and E. S. Lindstrom. 1971. Isolation and counting of Athiorhodaceae with membrane filters. Appl. Microbiol. 22:683–687. Tabita, F. R. 1995. The biochemistry and metabolic regulation of carbon metabolism and CO2 fixation in purple bacteria. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 885–914. Thiemann, B., and J. F. Imhoff. 1996. Differentiation of Ectothiorhodospiraceae based on their fatty acid composition. Syst. Appl. Microbiol. 19:223–230. Trüper, H. G. 1970. Culture and isolation of phototrophic sulfur bacteria from the marine environment. Helgol. Wiss. Meeresunters. 20:6–16. Uffen, R. L. 1973. Growth properties of Rhodospirillum rubrum mutants and fermentation of pyruvate in anaerobic, dark conditions. J. Bacteriol. 116:874–884. Uffen, R. L. 1978. Fermentative metabolism and growth of photosynthetic bacteria. In: R. K. Clayton, and W. R. Sistrom (Eds.) The Photosynthetic Bacteria. Plenum Press. New York, NY. 857–872. Uffen, R. L. 1983. Metabolism of carbon monoxide by Rhodopseudomonas gelatinosa: Cell growth and properties of the oxidation system. J. Bacteriol. 155:956– 965. Van Niel, C. B. 1944. The culture, general physiology, morphology and classification of the nonsulfur purple and brown bacteria. Bacteriol. Rev. 8:1–118.

64

J.F. Imhoff

Van Niel, C. B. 1971. Techniques for the enrichment, isolation, and maintenance of photosynthetic bacteria. In: S. P. Collowick and N. V. Kaplan (Eds.) Methods in Enzymology, Volume 23, Part A. Academic Press. New York, NY. 3–28. Vincenzini, M., R. Materassi, M. R. Tredici, and G. Florenzano. 1982. Hydrogen production by immobilized cells. I: Light dependent assimilation of organic substance by Rhodopseudomonas palustris. Int. J. Hydrogen Energy 7:231–236. Von Felten, P., H. Zürrer, and R. Bachofen. 1985. Production of molecular hydrogen with immobilized cells of Rhodospirillum rubrum. Appl. Microbiol. Biotechnol. 23:15–20. Vrati, S., and J. Verma. 1983. Production of molecular hydrogen and single cell protein by Rhodopseudomonas capsulata from cow dung. J. Ferm. Technol. 61:157–162. Vrati, S. 1984. Single cell protein production by photosynthetic bacteria grown on the clarified effluents of a biogas plant. Appl. Microbiol. Biotechnol. 19:199–202. Weckesser, J., H. Mayer, and G. Schulz. 1995. Anoxygenic phototrophic bacteria: Model organisms for studies on cell wall macromolecules. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 207–230. Weetall, H. H., B. P. Sharma, and C. C. Detar. 1981. Photometabolic production of hydrogen from organic substrates by free and immobilized mixed cultures of Rhodospirillum rubrum and Klebsiella pneumoniae. Biotechnol. Bioeng. 23:605–615. Westmacott, D., and S. B. Primrose. 1975. An anaerobic bag for photoheterotrophic growth of some Rhodospirillaceae in petri dishes. J. Applied Bacteriol. 38:205–207. Winogradsky, S. 1888. Beiträge zur Morphologie und Physiologie der Bakterien, Volume 1: Zur Morphologie und Physiologie der Schwefelbakterien. Verlag A. Felix. Leipzig, Germany.

CHAPTER 3.1.1 Woese, C. R., E. Stackebrandt, W. G. Weisburg, B. J. Paster, M. T. Madigan, V. J. Fowler, C. M. Hahn, P. Blanz, R. Gupta, K. H. Nealson, and G. E. Fox. 1984a. The phylogeny of purple bacteria: The alpha subdivision. Syst. Appl. Microbiol. 5:315–326. Woese, C. R., W. G. Weisburg, B. J. Paster, C. M. Hahn, R. S. Tanner, N. R. Krieg, H.-P. Koops, H. Harms, and E. Stackebrandt. 1984b. The phylogeny of purple bacteria: The beta subdivision. Syst. Appl. Microbiol. 5:327–336. Woese, C. R., W. G. Weisburg, C. M. Hahn, B. J. Paster, L. B. Zablen, B. J. Lewis, T. J. Macke, W. Ludwig, and E. Stackebrandt. 1985. The phylogeny of purple bacteria: The gamma subdivision. Syst. Appl. Microbiol. 6:25–33. Woese, C. R. 1987. Bacterial evolution. Microbiol. Rev. 51:221–271. Xia, Y., T. M. Embley, and A. G. O’Donnell. 1994. Phylogenetic analysis of Azospirillum by direct sequencing of PCR amplified 16S rDNA. Syst. Appl. Microbiol. 17:197–201. Yen, H.-C., and B. Marrs. 1977. Growth of Rhodopseudomonas capsulata under anaerobic dark conditions with dimethyl sulfoxide. Arch. Biochem. Biophys. 181:411– 418. Yildiz, F. H., H. Gest, and C. E. Bauer. 1991. Attenuated effect of oxygen on photopigment synthesis in Rhodospirillum centenum. J. Bacteriol. 173:5502–5506. Zannoni, D. 1995. Aerobic and anaerobic electron transport chains in anoxygenic phototrophic bacteria. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 949–971. Zuber, H., and R. J. Cogdell. 1995. Structure and organization of purple bacterial antenna complexes. In: R. E. Blankenship, M. T. Madigan, and C. E. Bauer (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 315–348.

Prokaryotes (2006) 5:65–71 DOI: 10.1007/0-387-30745-1_3

CHAPTER 3.1.2 ehT

areneG

mu i bor c imoceht sorP

dna

mu i bor c imo l acnA

The Genera Prosthecomicrobium and Ancalomicrobium GARY E. OERTLI, CHERYL JENKINS, NAOMI WARD, FRED RAINEY, ERKO STACKEBRANT AND JAMES T. STALEY

The genera Prosthecomicrobium and Ancalomicrobium are heterotrophic prosthecate bacteria, which produce numerous prosthecae per cell extending in all directions. These prosthecae vary in size and number both within and among species. Morphologically Prosthecomicrobium and Ancalomicrobium resemble one another because of their budding division and multiple prosthecae, but the former group is obligately aerobic whereas Ancalomicrobium strains are facultative anaerobes. Images of the type species Prosthecomicrobium pneumaticum and Ancalomicrobium adetum are shown in Figs. 1 and 2, respectively.

Taxonomy Phylogenetic analysis of the 16S ribosomal RNA gene sequences of P. pneumaticum, P. enhydrum, P. hirschii, A. adetum, and several uncharacterized Prosthecomicrobium-like and Ancalomicrobium strains demonstrate that these organisms belong to the Alphaproteobacteria (Fig. 3). This group contains other budding and prosthecate bacteria, including representatives of the bacterial genera Rhodomicrobium and Rhodopseudomonas as well as Hyphomicrobium and Caulobacter. These findings are consistent with earlier 16S rRNA cataloguing studies on these genera (Schlesner et al., 1989). Ancalomicrobium comprises a coherent clade within this group. Notably, Ancalomicrobium carries out a mixed-acid type fermentation, the products of which are identical to those of Escherichia coli, a member of the Gammaproteobacteria (Van Neerven and Staley, 1988). Ancalomicrobium is the only member of the Alphaproteobacteria currently known to have a mixed-acid fermentation. Analyses of the 16S rRNA genes demonstrate that the genus Prosthecomicrobium is polyphyletic (Fig. 3) as none of the type strains of the other two species clusters with the type species, P. pneumaticum. Three strains, including P. enhydrum (ATCC 23634) and two Prosthecomicrobium-like bacteria, AP4.6 and P3.12, cluster with

Devosia neptuniae (LGM 21357), a nonprosthecate organism. SCH75 is a morphologically distinctive organism whose multi-appendaged cells produce a holdfast from one polar prostheca (Schlesner et al., 1989). This representative, along with SCH127, is more closely related to members of the Mesorhizobium genus than to other members of Prosthecomicrobium. Prosthecomicrobium hirschii (ATCC 27832) and SCH71 form deep branches unrelated to any named genera. Finally SCH235 clusters with members of several named genera. This analysis suggests that the genus Prosthecomicrobium should be revised taxonomically. Chemical analysis of phospolipids from different Prosthecomicrobium strains indicated that there are at least five distinct groups (Sittig and Schlesner, 1993). Four of these groups are represented in Fig. 3, with members of each group clustering in distinct phylogenetic positions. Additionally, various species and several as yet unnamed strains show little or no intrageneric DNA-DNA hybridization with one another (Moore and Staley, 1976; Schlesner et al., 1989; Chernykh et al., 1990). To our knowledge, 16S rRNA sequencing of the remaining characterized species (Table 1) P. litoralum (Bauld et al., 1983), “P. polyspheroidum,” “P. consociatum,” and “P. mishustinii,” (Vasilyeva and Lafitskaya, 1976; Vasileva et al., 1991) has not yet been reported and will be important in assessing the extent of diversity within the Prosthecomicrobium genus. Reclassification of the bacteria that have been analyzed is required, as they do not cluster with the type species of Prosthecomicrobium, P. pneumaticum. The named species should therefore be reassigned to different genera.

Habitats Multi-appendaged bacteria have been reported from a variety of natural habitats. Many strains have been isolated from fresh water (Staley, 1968), brackish and marine water (Bauld et al., 1983; Schlesner et al., 1989), groundwater (Hirsch

66

G.E. Oertli et al.

CHAPTER 3.1.2

and Rades-Rohkohl, 1983), soil (Vasil’eva et al., 1974), and pulp mill aeration ponds (Stanley et al., 1979), where they may occur in high concentrations (Fig. 4). In one report (Bianchi, 1989), what appears to be a species of Ancalomicrobium was unexpectedly found in high concentrations in seawater used for shrimp aquaculture. Apparently the long prosthecae make these bacteria resistant to grazing by protozoan predators. Bacteria with multiple appendages have also been reported in the intestinal tracts of insects (Cruden and Markovetz, 1981), possibly because of concentration by feeding. The multi-appendaged prosthecate bacteria appear to be well adapted for growth in olig-

otrophic and mesotrophic soil and aquatic habitats. Chemostat studies of Prosthecomicrobium hirschii indicate that their maximum growth rates are low regardless of the concentration of carbon source (Semenov and Vasileva, 1986). However, they have a high affinity for certain substrates such as glucose, as well as a low rate of respiration. These data suggest that in environments with low sugar concentrations these bacteria can metabolize efficiently because they have high-affinity uptake systems, which provide them with their selective advantage. Thus, their slow growth and low respiration rates may be advantages in oligotrophic environments where they reside (Semenov and Vasileva, 1986; Semenov and Staley, 1993). Autoradiographic studies of acetate uptake in pulp mill aeration lagoons by Ancalomicrobium indicated that Ancalomicrobium was not as important as other bacteria in the metabolism of this substrate (Stanley and Staley, 1977). While their in situ uptake of sugars, which are abundant in this habitat, was not examined, it is clear that they are metabolically active in this environment.

Enrichment and Isolation

Fig. 1. Prosthecomicrobium pneumaticum (ATCC 23633). Transmission electron micrograph showing gas vesicles within the cell, many short prosthecae, and a single long prostheca. Bar = 1.0 µm.

A

B

These bacteria occur in relatively low concentrations in natural habitats, typically 0.1–1 cell per ml (Staley et al., 1980), although they have been found in concentrations greater than 106 cells per ml in pulp mill aeration lagoons (Stanley et al., 1979). Generally, prior to isolation of pure cultures it is necessary to enrich for them. The usual procedure for enrichment entails using lownutrient media. The most common procedure is to add a sterile peptone solution to a natural water sample to give a final concentration of 0.01% (Houwink, 1951). Schlesner et al. (1989) used a variety of carbohydrates (0.05% glucose,

Fig. 2. Ancalomicrobium adetum (ATCC 23632). A) Phase contrast image of A. adetum cells showing long appendages and including one with a branch and B) scanning electron micrograph. Bar = 5.0 µm.

CHAPTER 3.1.2 Fig. 3. Phylogeny of 16S rRNA genes from various Prosthecomicrobium and Ancalomicrobium isolates. Rooted neighbor-joining tree showing evolutionary relationships between various Ancalomicrobium and Prosthecomicrobium strains (bold). Reference sequences were picked by comparing these to 92,813 rRNA sequences in the RDP-II database using the sequence match tool (Cole et al., 2003), and complete sequences with the highest similarity to the query were used in our tree. Distances were calculated using the Kimura-2parameter test over a total of 1137 characters. Escherichia coli was used as an outgroup. Percentages at nodes, with values greater than 50% shown, indicate bootstrap values for 1000 replications of the dataset; a-d indicate groups classified by their phospholipid composition as examined by Sittig and Schlesner (1993). a—group 1; b— group 2; c—group 3; and d— group 5. Bar represents the 10% 16S rDNA sequence distance.

The Genera Prosthecomicrobium and Ancalomicrobium

67

Prosthecomicrobium pneumaticumb Ancalomicrobium sp. A2.22 Ancalomicrobium sp. A2.7 100 Ancalomicrobium sp. A18 Ancalomicrobium sp. 3.15 100 Ancalomicrobium adetum Proteobacterium F0723 57 52 100 Proteobacterium 34619 Prosthecomicrobium hirschii strain 16d Angulomicrobium amantiformis 86 Ancylobacter rudongensis 100 100 Starkeya novella 52 82 Prosthecomicrobium sp. SCH235; IFAM3000 80 uncultured Proteobacterium SM1E02 71 93 Antarctic bacterium BR-10753a 74 Devosia neptuniae 95 uncultured Proteobacterium Blci23 Prosthecomicrobium enhydruma 99 Prosthecomicrobium sp. P3.12 88 100 Prosthecomicrobium sp. AP4.6 uncultured Proteobacterium Blri24 60 Prosthecomicrobium sp. SCH300; IFAM3210 100 Rhizobium leguminosarum 86 100 Rhizobium etli 94 Ochrobactrum tritici Prosthecomicrobium sp. SCH127; IFAM1314c 66 Prosthecomicrobium sp. SCH75; IFAM1508 98 Mesorhizobium loti 95 Mesorhizobium tianshanense 99 99 Mesorhizobium mediterraneum uncultured sludge bacterium A20 uncultured bacterium Y14-4a 100 71SCH; IFAM1551 100 Stella vacuolata Escherichie coli 0.1

Table 1. Species of Ancalomicrobium and Prosthecomicrobium.a Species name

Type strain

Source

Ancalomicrobium adetum Prosthecomicrobium pneumaticum Prosthecomicrobium enhydrum Prosthecomicrobium hirschii Prosthecomicrobium litoralum

4a (ATCC 23632) 3a (ATCC 23633) 9b (ATCC 23634) 16 (ATCC 27832) 524-16 (ATCC 35022)

“Prosthecomicrobium mishustinii” “Prosthecomicrobium consociatum” “Prosthecomicrobium polyspheroidum”

17 11

Putah Creek, CA, USA Putah Creek, CA, USA Putah Creek, CA, USA Pond, NC, USA Seawater, Puget Sound, USA Soil, Colchis, Russia Compost, Shermetovo, Russia Soil, Russia

a

Although the species in quotation marks are in culture, their names have not been validly published.

or 0.01% lactate, or 0.1% glucosamine) or 0.25% CaCO3 or 0.1% NaNO3 in their enrichments. A typical enrichment procedure is as follows: 10 mg of peptone is added to a 150-ml graduated beaker, which is covered with aluminum foil and autoclaved. Alternatively, a volume of an autoclaved peptone solution, equivalent to 10 mg, can be added to the beaker. A fresh- or marine-water sample is collected aseptically in a sterile bottle

and 100 ml is transferred to one of the prepared beakers. This is then incubated at room temperature for 1–4 weeks, during which wet mounts are examined periodically using a phase microscope. When numbers of these organisms become appreciable, they may be isolated by streaking portions or spread-plating dilutions (10–3 to 10–5) on dilute peptone agar (DPA). Additionally, these bacteria can be isolated from

68

G.E. Oertli et al.

CHAPTER 3.1.2 Nicotinamide Pyridoxine HCl Riboflavin Thiamine HCl

5 mg 10 mg 5 mg 5 mg

Add distilled water up to 1 liter and store at 4°C in a dark container.

Fig. 4. Electron micrograph of a pulp mill environmental sample showing a high concentration of prosthecate bacteria.

The DPA plates should be incubated for about two weeks at room temperature prior to examination. Colonies are best located using a binocular dissecting microscope. The most likely colonies are small, have an entire margin, and may be a variety of colors (Table 1). Wet mounts of prospective colonies should be examined with a phase microscope, and when a presumptive colony has been located, it should be restreaked for purification. If the entire colony has been used in preparation of the wet mount, a loopful of the wet mount can be removed and used for streaking. This material may be streaked on DPA again, although a richer medium, MMB (Modified Medium B), will permit more rapid growth. MMB Agar

liquid enrichment cultures by passing them through a column containing glass beads and plating eluted fractions (Staley, 1968). Dilute Peptone Agar (DPA) Peptone Hutner’s modified salts solution (see below) Vitamin solution (see below) Agar

0.1 g 20 ml 10 ml 15 g

Add distilled water up to a total of 1 liter.

Hutner’s Modified Salts Solution Nitrilotriacetic acid MgSO4 · 7H2O CaCl2 · 2H2O NaMoO4 · 2H2O FeSO4 · 7H2O Metals “44”

10.0 g 29.7 g 3.3 g 12.7 mg 99.0 mg 50 ml

First neutralize the nitrilotriacetic acid with potassium hydroxide, then add the remaining ingredients. Adjust the pH to 7.2 with KOH and H2SO4. Add distilled water up to a total of 1 liter. Metals “44” contains per 100 ml: ethylene diamine tetraacetic acid, 250 mg; ZnSO4 · 7H2O, 1095 mg; FeSO4 · 7H2O, 500 mg; MnSO4 · H2O, 154 mg; CuSO4 · 5H2O, 39.2 mg; CoCl2 · 6H2O, 20.3 mg; and Na2B4O7 · 10H2O, 17.7 mg. Add a few drops of sulfuric acid to prevent precipitation before making to volume with distilled water, and store at 4°C. Vitamin Solution B12 Biotin Calcium pantothenate Folic acid

0.1 mg 2 mg 5 mg 2 mg

Peptone Yeast extract Glucose Ammonium sulfate Hutner’s modified salts solution Vitamin solution Agar

0.15 g 0.15 g 1.0 g 0.25 g 20 ml 10 ml 15 g

Add distilled water up to 1 liter. Adjust pH to 7.0–7.5 prior to autoclaving.

In habitats where their concentrations relative to other bacteria are much higher, isolation may be effected by streaking directly onto MMB agar. This practice has been used to isolate strains from pulp mill oxidation pond samples.

Cultivation and Maintenance For routine cultivation and maintenance, cultures may be grown on MMB agar. Some strains tested seem to grow well on the richer and more easily prepared medium used for cultivation of caulobacters (Poindexter, 1964). This medium contains 0.2% peptone, 0.1% yeast extract, 0.02% magnesium sulfate, and 1.5% agar and is made to volume with tap water. Though higher yields may be obtained with this medium, pleomorphic forms occur with some strains. For example, Ancalomicrobium adetum produces high yields on this medium, but cells may form short chains and mini cells with one or two prosthecae. Alternatively a more concentrated version of MMB, termed “Super” MMB, has been found to work well for cultivation of Ancalomicrobium strains (Van Neerven and Staley, 1988). These bacteria can also be grown on a defined medium containing ammonium sulfate as sole

CHAPTER 3.1.2

The Genera Prosthecomicrobium and Ancalomicrobium

nitrogen source and glucose as sole carbon source. The composition of this medium is as follows: Defined Medium Ammonium sulfate Disodium phosphate Glucose Hutner’s modified salts solution Vitamin solution

0.25 g 0.0005 M 0.25 g 20 ml 10 ml

Add distilled water up to a total of 1 liter. For solid medium, add 1.5% agar.

This medium has been useful in ascertaining vitamin requirements using the single-deletion method (Staley, 1968). It has also been of limited use in determining carbon source utilization patterns, although MMB from which glucose has been deleted is generally more useful for that purpose. Slant cultures maintain viability for at least one month at refrigerator temperatures. Lyophilization is satisfactory for longer periods of preservation. Frozen cultures made using 15% (w/v) glycerol and stored at –70°C have proven to be viable for at least 4 years. Storage of glycerol stocks at –20°C is not recommended.

Identification Table 2 shows some of the major differences among the four validated and two nonvalidated species of Prosthecomicrobium and one species of Ancalomicrobium that have been described at this time.

69

The first step in determining whether an unidentified bacterium is a representative of one of these genera involves examination of a wet mount of a suspected colony under a phase microscope using an oil-immersion objective. Prosthecae on most strains can be detected by ordinary phase microscopy, but if there is any uncertainty, whole cells of the suspect strain must be examined using a transmission electron microscope (TEM). Most species of Prosthecomicrobium produce relatively short appendages (i.e., 500 0.003 20 0.15 0.002 yes 20 (strong) 0.2 2

Cherwonogrodzky et al., 1990 Cherwonogrodzky et al., 1990 Cherwonogrodzky et al., 1990 Cherwonogrodzky et al., 1990 Cherwonogrodzky et al., 1990 Cherwonogrodzky et al., 1990 Cherwonogrodzky et al., 1990 Rasool et al., 1992 Rasool et al., 1992

200 ≥10 ≥10

50 0.007 0.023

Goldstein et al., 1992 Goldstein et al., 1992 Goldstein et al., 1992

Abbreviations: LPS, lipopolysaccharide(s); C′, complement; IFN, interferon; IL, interleukin; and TNF, tumor necrosis factor. µg causing the indicated effect. b Carrageenan-treated endotoxin-resistant mice. a

2000

Enterobacterial LPS / Brucella LPS

1500

1000

500

e

b oz

ym

-1 Ly s

IL

Fa TN

ci To xi

re TB N

LPS Biological Activities Lipid A is the structure directly involved in the endotoxin-related properties of LPS. Since the lipid A of Brucella has a structure that clearly departs from that of the classical enterobacterial-type lipid A, it is not surprising that its LPS shows lower biological activities (Table 14). In general, the lower endotoxicity of Brucella LPS is manifested as a marked reduction in toxicity linked to a reduced ability to stimulate susceptible host cells to release immune mediators. Indeed, Brucella LPS is from 200 to 2,000 times less active than Salmonella LPS (Fig. 17). Brucella abortus LPS activates the complement system poorly by the alternative pathway, and the O-polysaccharide has been shown to be involved in this phenomenon (Fig. 18). Both reduced endotoxicity and poor complement activation contribute to virulence (Opsonization and Complement Susceptibility). There is no evidence of interaction between the O-polysaccharide of B. abortus and collectins (mannose-binding protein and surfac-

ty

0

d

netically distant bacteria (Table 6). Since the genetic and ecological characteristics of the brucellae suggest that they presently have little opportunities for such exchanges (Genetic Exchange, Plasmids and Lysogenic Phages), acquisition of O-polysaccharide genes could have been an early event in the evolution of the genus (Acquisition of Ancestral Genes by Horizontal Transference). This is in keeping with the structural homogeneity of the N-formylperosamine homopolymers and with the presence in B. ovis and B. canis of genes involved in its synthesis (Cloeckaert et al., 2000).

Fig. 17. Comparison of the biological activities of Brucella LPS with S. typhimurium and E. coli LPSs in phagocytic cells. The concentration of Brucella LPS at which the maximum biological effect is generated was recorded and compared with the concentration of S. typhimurium or E. coli for the same effect. Respiratory shunt in human polymorphonuclear leukocytes was measured as reduction of nitro-blue tetrazolium (NTB-red); maximum level of toxicity (Toxicity) reached with B. abortus LPS was less that 10% at 500 mg/ml × 106 macrophages; TNFα production (TNF-α) in murine macrophages; IL-1β production (IL-1β) by murine macrophages; and lysozyme release (Lysozyme) by human polymorphonuclear leukocytes. Notice that Brucella LPS is from 200 to 2,000 times less potent than Salmonella or Escherichia LPSs. Adapted from Rasool et al. (1992); Goldstein et al. (1992).

356

E. Moreno and I. Moriyó n

CHAPTER 3.1.16

tant proteins), although B. abortus has been shown to bind to human B cells and macrophages by a lectin-type mechanism (Brucella Bacterins and Lipopolysaccharide as Immunological Tools). Native Hapten Polysaccharides These are haptenic polysaccharides that can be demonstrated in extracts of smooth Brucella by immuno-

Anticomplementary activity

100

E. coli Lipid A

PS

iL

75

E.

l Co

50 Brucella R-LPS

25 Brucella S-LPS

0 0

20

40

60

LPS mg Fig. 18. Anticomplementary activity of LPS and lipid A. Increasing amounts of each preparation were incubated with guinea pig complement and percentage loss of hemolytic activity was measured. Notice that B. abortus smooth and rough LPSs exert low anticomplementary activity only at high concentrations of material, whereas E. coli lipid A and Salmonella LPS display a powerful anticomplementary activity. Adapted from Moreno et al. (1981), with permission.

precipitation with sera from B. abortus- or B. melitensis-infected animals in the presence of hypertonic buffers (Fig. 19). The B. melitensis 115 rough mutant is unable to produce smooth-LPS but also accumulates a similar compound (named “polysaccharide B”) in the cell envelope and its phenotype is close to that of LPS wzm mutants (Table 5). Both NH and polysaccharide B are N-formylperosamine homopolymers with a frequency of α-(1→2) and α-(1→3) linkages similar to that of the O-polysaccharide (Fig. 16) of the strain from which they are isolated (Aragó n et al., 1996b). However, in contrast to the O-polysaccharide of LPS, neither polysaccharide B nor NH is linked to core oligosaccharide sugars and both are partially (up to 50%) deformylated (M. Staaf et al., unpublished observations). This last feature is intriguing because polymerization of at least B. melitensis LPS O-polysaccharides is blocked by disruption of the gene coding for the putative formyltransferase (Godfroid et al., 2000; Table 5). NH and polysaccharide B may represent biosynthetic precursors of the O-polysaccharide of smooth-LPS, but NH also has been demonstrated in the outer-membrane fragments that are released spontaneously by growing B. melitensis. Thus, NH is in the outer-membrane as a structure that, although similar to the Opolysaccharides, is not linked to core-lipid A (Outer Membrane Topology). This type of polysaccharide was described long ago in several Gram-negative bacteria (Anacker et al., 1964) and it is known as their presence in the outermembrane results from variations in the standard polymer-lipid A-core ligase reaction (Whitfield et al., 1997).

1 Poly B

A 2

NH

3 NH

B

LPS LPS

I I

V B

2

I

V 3 NH

B

I

LPS 4

Fig. 19. Immunoprecipitation analysis of Brucella polysaccharides and lipopolysaccharides. Left panel: immunoelectrophoresis of rough B. melitensis 115 polysaccharide B (A); and B. melitensis extracts containing native hapten (NH) and smooth LPS (B). The troughs contained sera from infected cattle either plain (1 and 3) or absorbed with smooth Brucella cells (2) or polysaccharide B (4). Right panel: Double gel immunodiffusion of B. melitensis extracts containing NH and smooth LPS and sera from infected (I) or B. abortus strain 19 vaccinated (V) cattle. Notice that while infected bovines react with both LPS and NH, vaccinated animals only react against the LPS, with no traces of reaction against the NH.

CHAPTER 3.1.16

The Genus Brucella

357

LPS

NH

NH-L

Braun’s Omp19

Lipid A

Kdo

Omp3a Omp10

OL

Omp2b

PC Omp19

OH-C28:0

Omp31 Braun’s -linked

Omp10

Omp3b

Omp16.5

Fig. 20. A hypothetical model of the Brucella outer membrane. Omps are indicated by their mass in kDa, except for two group 3 proteins (Omp3a and Omp3b) with molecular weights of 26 kDa and 23 kDa, respectively. Lipoproteins have their acyl groups inserted in the bilayer. The lipid A diaminoglucose disaccharide backbone (represented by yellow trapezoids) is linked to fatty acids (black zigzag structures). The red squares in the bottom of the structure mark the hydroxyl groups of the LPS acyl chains that span the outer membrane. Purple squares mark the reducing ends of the NHs, an unknown sugar that in some cases may be linked to a lipid (NH-L). Braun’s lipoprotein may be linked to peptidoglycan or free in the outer membrane. The polar heads of ornithine lipids (OL) are marked as green rectangles. Dark circles represent phospholipids, mainly phosphatidylcholine (PC). Notice that the NH of various sizes are intertwined with the O-polysaccharides of the LPS in the outer membrane, forming a dense layer. The negative charges of the LPS are neutralized by the positive charges of the ornithine lipids (OL) and cationic Omp such as Omp3b. Group 3 proteins (Omp3a [Omp25], Omp3b and Omp31) are strongly bound to LPS. Omp2b is a porin.

Outer Membrane Proteins The major Omps of Brucella were first identified by detergent extraction of cell envelopes and classified according to their apparent molecular weights as group 1 (94 or 88 kDa), group 2 (36–38 kDa), and group 3 (31–34 kDa and 25–27 kDa). Moreover, a lipoprotein linked to the peptidoglycan and three other lipoproteins have been identified in the cell envelope. Group 2 are porins and group 3 contains at least three unrelated proteins (Omp31, Omp3a or Omp25, and Omp3b), but the nature and role of group 1 remains to be elucidated (Fig. 20). Presently, little is known about the Omps of the Brucella isolated from marine mammals, and the data summarized in the following sections concern only the six classical species. Most information concerning the Brucella Omps has been reviewed by Cloeckaert et al. (1996b). Porin Proteins The brucellae carry two related porin genes (omp2a and omp2b) with a high

degree (>85%) of internal homology but with little homology to the genes of other porins described so far. This omp2 locus is present in all Brucella species and is highly polymorphic at both omp2a and omp2b. This locus contains some species-specific sequences, specific markers for several of the classical biovars, and other sorts of intraspecific diversity (Fig. 4), including natural Omp2a-Omp2b chimeric proteins (Paquet et al., 2001). Thus, sequence analysis of the omp2 genes has also taxonomical and phylogenetic implications (Table 4). Most of these variations are created by simple exchanges of conserved motifs between omp2a and omp2b and, in general, seem to have little impact on the antigenic variability of the proteins. The cognate proteins are predicted to be folded as a 16 stranded β-barrel, and most differences seem accumulated at the external loops (Paquet et al., 2001). At least Omp2b is in a trimeric state in the outer- membrane and behaves as a peptidoglycan-associated protein (Fig. 20). Obvi-

358

E. Moreno and I. Moriyó n

ously, Brucella porins span the outer-membrane and contribute to its permeability. In vitro, B. abortus only expresses Omp2a, and Omp2b may be expressed under other conditions. The pattern of expression in other Brucella species is not known. Omp31 The omp31 gene is present and expressed in all Brucella classical species but B. abortus, which carries a long 8-kb deletion encompassing this gene and others of unknown function (Vizcaíno et al., 1997). Gene omp31 shows only little homology with omp3a (omp25) or omp3b, the other two groups of genes coding for a group 3 protein. Omp31 shows a tight interaction with LPS and is associated with peptidoglycan (Outer Membrane Topology). These features suggest that this protein plays a structural role, but if so, its absence from B. abortus suggests that it is not essential. Omp31 is able to form oligomers resistant to denaturation, and since this is a property characteristic of some porins, it has been suggested that it may play this role (Fig. 20). Omp3a and Omp3b Family Two different groups of Omps, Omp3a (formerly Omp25) and Omp3b, are composed of 8 beta sheets and are related to RopA and RopB proteins of Rhizobium (Fig. 20). These two protein families have a molecular weight ranging from 26–23 kDa and each group displays from 5 to 8 spots in two dimensional gels (C. Guzmán-Verri et al., unpublished results). Several putative genes coding for these proteins have been identified in the Brucella genome, supporting the two-dimensional gel findings and supporting that they constitute a family of Omps with unknown functions. Similarly to Omp31, they show characteristically tight interaction with the LPS and are also associated with the peptidoglycan (Outer Membrane Topology). Omp3a is highly conserved in Brucella species and the only remarkable variation is a 36-bp deletion at the 3’ end of the B. ovis omp3a gene that causes an antigenic shift in the Omp3a of this species. Although both families may play a structural role, this does not exclude additional functions in virulence, since at least in B. abortus, its expression in the outer-membrane seems to be regulated by the BvrR-BvrS system (Mechanisms of Entry to Host Cells). Lipoproteins The Brucella cell envelope contains three proteins (Omp19, Omp16.5 and Omp10) with genetic features characteristic of bacterial lipoproteins and which have been shown to incorporate palmitic acid (Tibor et al., 1999). These lipoproteins are similar to the peptidoglycan-associated lipoproteins of other Gram-negative bacteria (Outer Membrane

CHAPTER 3.1.16

Topology) and crossreact with homologous proteins of other members of the α-2 Proteobacteria. An additional lipoprotein of overall characteristics similar to those of Braun’s lipoprotein has been shown in B. abortus, B. melitensis and B. ovis (Gó mez-Miguel and Moriyó n, 1986), but the corresponding gene has not been identified (Fig. 20). Outer Membrane Topology Like in other Gram-negative bacteria, the LPS is located in the outer leaflet of the outer-membrane anchored by its lipid moiety (Fig. 20). The very long fatty acids of the lipid A are likely to span the outermembrane and to be anchored on the inner surface by the hydroxyl group close to the end of the acyl chain. The phosphate groups in lipid A and the Kdo carboxyl groups of the core may be associated with ornithine lipids, because the latter lipids appear as contaminants in LPS extracts and have a free amino group with a positive charge (Freer et al., 1996). With regard to the phospholipids, there is evidence that phosphatidylcholine is preferentially distributed in the outer-membrane (Gamazo and Moriyó n, 1987), perhaps in its inner leaflet. In the smooth strains, the O-polysaccharide extends outwards, in association with the NH. This last molecule is maintained in the outer-membrane intertwined with the LPS O-polysaccharide or linked to a lipid other than lipid A (Rojas et al., 1994). The presence or the absence of the O-polysaccharide profoundly modifies the topology of the surface of Brucella cells. First, it masks the ionic groups of the proteins and the core-lipid A LPS sections. Since the latter sections are dominant, mutants lacking the O-polysaccharide have a negatively charged surface at physiological pH, which contrasts with the very weak charge carried by smooth cells (Schurig et al., 1981; Weber et al., 1978). This difference is the basis of some tests used to distinguish smooth strains from their rough mutants (Cellular and Colonial Characteristics). Second, access to the anionic groups of the lipid A-core is sterically hindered in the smooth strains and this contributes to resistance to polycations (Properties of the Outer Membrane). Finally, there are differences in the surface exposure of Omps between the smooth Brucella species on the one hand and the rough mutants and B. ovis and B. canis on the other. In general, these differences are manifested as a comparatively reduced Omp accessibility to antibodies in the smooth strains. However, not all smooth strains are identical in this property, and the variations reflect at least in part different degrees of O-polysaccharide polymerization and also the total amount of smooth LPS produced. In some smooth strains studied in detail, Omp accessibility to antibodies is par-

CHAPTER 3.1.16 Binding to cells

The Genus Brucella

Nfpp

Virulence for host

B. abortus 45/20 B. abortus 9.49 (per A– ) B. melitensis B115 B. abortus RB51

B. abortus BvrS B. abortus BvrR B. abortus BvrS-R

B. abortus S19 B. ovis Reo 198 and field isolate B. canis RM6/66 and field isolate B. suis 503, 1330, IB-2579 B. abortus 2308 and 544 B. melitensis 16M B. neotomae 418

Fig. 21. Correlation among Brucella strains binding to epithelial cells, presence of N-formylperosamine polysaccharides (Nfpp) and virulence. Values are ranked from negative or absent (−), weak positive or present in low quantities r (+) to strong positive or present in large quantities (++++). The base of the triangle is compatible with the high number of bacteria binding to cells, whereas the apex of the triangle is compatible with the low number of brucellae binding to cells. Notice the absence of correlation between presence of Detilleux et al. (1990a, b); Freer et al. (1999); and PizarroCerdá (1998).

alleled by an increase in both sensitivity to the R/C (rough-specific) Brucella phages (The Brucellaphages) and exposure of LPS core-lipid A epitopes. However, not all rough mutant strains are identical in Omp antibody accessibility, and differences between B. ovis and B. canis also have been observed (Bowden et al., 1995). Interestingly, these differences correlate well with the differences observed between the “natural rough” and rough mutants in the attachment and invasion of cells (Fig. 21). Several Omps behave as peptidoglycanassociated proteins. If molecular weight, surface exposure and association with the peptidoglycan are used as criteria, porins, Omp31 and Omp3a are membrane-spanning proteins. The topology of Omp19, Omp16.5 and Omp10 proteins is probably more complex. They are accessible to antibodies on the surface of rough strains, and although according to the predicted amino acid sequence are hydrophilic, they also carry a lipid moiety and are extracted with detergents. Their similarities with the peptidoglycan-associated lipoproteins of other Proteobacteria suggest that they could be in the periplasmic space and play a role in the interaction of the outer-membrane with the peptidoglycan but they should also be anchored on the cell surface. The Braun’s lipoprotein of Brucella seems to be covalently attached to the peptidoglycan but it also has

359

been reported to be surface exposed, perhaps as a free form (Fig. 20). Properties of the Outer Membrane Some simple observations, such as the inability to grow in media containing hydrophobic dyes, the ability to grow on selective media containing polymyxin B (Table 11), and the acid-fastness when stained by Stamp’s method (Fig. 10) (Cellular and Colonial Characteristics), show that the brucellae have cell envelopes with a characteristic set of properties, some of which reflect, in all likelihood, an adaptation to intracellular parasitism (Outer Membrane Versus Bactericidal Substances). Permeability Nutrients or precursors must passively cross the outer-membrane before they are taken up by the periplasmic and membrane proteins engaged in transport and translocation through the cytoplasmic membrane. Some early observations on the oxidation of glutamate by resting cells suggest that there is a link between outer-membrane permeability and virulence (Oxidation of Amino Acids). This could occur as a result of subtle differences in outer-membrane topology indirectly causing alterations in the pathways described (The Hydrophilic Pathway). There are two possible pathways for substrates to penetrate through Gram-negative outermembranes and both are accessible in Brucella cells. The Hydrophilic Pathway Some small hydrophilic molecules penetrate the outer-membrane of Brucella through Omp2 porins (Douglas et al., 1984). In classical functional assays, porins of B. abortus, B. melitensis and B. canis show an apparent internal diameter of about 1.2 nm, comparable to that of the OmpF porin of E. coli. At least for B. abortus, this result should correspond to Omp2b, since Omp2a is not expressed in vitro. However, cloned B. abortus Omp2a increases the outer-membrane permeability of E. coli to maltodextrins, and in planar bilayers, Omp2a forms monomeric pores smaller than those of Omp2b (the latter shows channel activity in trimeric state) (Marquis and Ficht, 1993). Although only three species and a few representative strains have been examined, there are small differences in the selectivity of porins purified from cell envelopes in functional assays according to which B. melitensis porin could have the smallest internal diameter and B. canis the largest, with B. abortus showing intermediate diameters (Douglas et al., 1984). Since thionin (a dye used in Brucella biotyping) has an exceptionally low hydrophobicity among dyes and its size is compatible with that of the Brucella porin channel, it has been suggested that differences in thionin

360

E. Moreno and I. Moriyó n

CHAPTER 3.1.16

sensitivity of B. melitensis and B. abortus (Table 3) relate to different functional properties of the porins (Douglas et al., 1984). Recent comparative analyses suggest functional differences between Omp2a and Omp2b, the former being more efficient in allowing sugar diffusion (Paquet et al., 2001). The Hydrophobic Pathway The antibiotic and dye sensitivity patterns and the direct assessment of permeability with hydrophobic probes show that the outer-membrane of Brucella is readily permeable to hydrophobic compounds (Martínez de Tejada and Moriyó n, 1993; Fig. 22). The same evidence suggests that this permeability is more marked in B. ovis than in B. abortus and B. melitensis (Freer et al., 1999), but comparative studies with a representative number of strains have not been performed. The well-known variations in sensitivity to the large hydrophobic dye basic fuchsin (Table 3) also suggest species and biovar differences in this pathway, which could reflect subtle outer-membrane structural or topological differences. This pathway also accounts for the effect of sexual hormones on Brucella cell envelopes and metabolic rates (Meyer, 1976) and for the inability of Brucella to grow on media designed for bacteria that, like Escherichia, Salmonella, Pseudomonas and many others, have efficient outer-membrane barriers to hydrophobic permeants. In Brucella, permeability to hydrophobic compounds is linked to the LPS structure (Freer et al., 1996), and comparative studies with Ochrobactrum sug-

Brucella abortus

Relative fluorescence

120

80

40 NPN

Escherichia coli

0 0

15

30 seconds

45

60

Fig. 22. Outer membrane permeability to the hydrophobic probe N-phenyl-1-naphthylamine (NPN). The arrow marks the time at which the bacteria were exposed to NPN and partition of the probe into the cell envelope (detected as a fluorescence increase) begins. Notice the increased uptake and permeability for the hydrophobic NPN probe by Brucella in comparison with the strong resistance to NPN penetration by E. coli cells. For details see Martinez-de-Tejada and Moriyó n (1993).

gest that the absence of negatively charged groups and counterions at core level (Fig. 6) plays an important role in the inability to establish an effective barrier (Velasco et al., 2000). Outer Membrane Stability It is a common observation that, as compared to E. coli, the brucellae are resistant to physical disintegration. Likewise, detergent extraction of Brucella cell envelope components needs conditions harsher than those used in many Gram-negative bacteria (reviewed in Moriyó n and Ló pez-Goñ i, 1998). Finally, Brucella cell envelopes are not destabilized by EDTA or polycationic microbicidal peptides and proteins that have a strong action on the majority of Gram-negative bacteria examined (Freer et al., 1996). At least in part, these observations are accounted for by the structure of the Brucella cell envelope. Omp-peptidoglycan Association Electron microscopy shows that the peptidoglycan of Brucella is in an interaction with the outermembrane much tighter than that observed in Escherichia coli (Fig. 12). Moreover, in some extraction procedures, Omps appear associated with the peptidoglycan. Although there are conflicting views on the precise structural meaning of this association (Moriyó n and Ló pez-Goñ i, 1998), it could provide the Brucella cell envelope with additional stability. Lipid Composition As compared to Escherichia, Salmonella and Pseudomonas, Brucella contains free lipids that carry longer acyl chains suggestive of a stronger hydrophobicity (Table 12). Moreover, phosphatidylcholine, which is the major phospholipid, has structural properties different from those of phosphatidylethanolamine (the major phospholipid in many other Gram-negative organisms), and at least in vitro, it forms more stable bilayers. Long acyl chains are also a characteristic of the Brucella lipid A (Fig. 7), and the overall hydrophobic effect could be reinforced by the topology of the very long fatty acids (Outer Membrane Topology). These features are present in other members of the α2 Proteobacteria with which the brucellae share the property of the acid-fastness when stained by the modified Stamp’s method (Fig. 10). Lipopolysaccharide and Resistance to Outermembrane Destabilizing Agents The reduced presence of anionic groups in the LPS core oligosaccharide and its strong hydrophobic anchorage (Fig. 20) are the main reasons for the marked resistance of Brucella to polymyxins (Fig. 23) and other polycationic peptides (Freer et al., 1996; Velasco et al., 2000; Fig. 24; Outer Membrane Versus Bactericidal Substances). Moreover,

CHAPTER 3.1.16

Ochrobactrum

361

Escherichia

Control

Brucella

Polymixin B

Fig. 23. Bactericidal activity of polymyxin B against B. abortus, O. intermedium and E. coli. Notice the characteristic outer membrane blebbing caused by polymyxin B in E. coli but not in B. abortus or O. intermedium. Brucella abortus treated with polymyxin B looks intact and not significantly different from the control. On the contrary, E. coli and O. intermedium are killed by polymyxin B. Although the outer membrane is morphologically unharmed, selfpromoted uptake of polymyxin B in O. intermedium is inferred from the extensive damage to the cytoplasmic membrane and from the subsequent cytoplasm coagulation. From Velasco et al. (2000), with permission.

The Genus Brucella

0.25 µm

% of bactericidal action 0

Fig. 24. Bactericidal activity of various cationic peptides and microbicidal substances against B. abortus and S. typhimurium. Concentrations used were the minimal quantity of bactericidal substance required to reduce the Salmonella colony-forming units (CFUs) not less than 75% in more than 99% of the controls. Incubation and number of organisms tested were the same for both bacteria: Notice the high resistance of Brucella to bactericidal substances in comparison to Salmonella. Adapted from Martínez de Tejada et al., 1995; Freer et al., 1996 and Páramo et al., 1998.

25

50

75

100

Bactenecin 7 Bactenecin 5 Cap18 Cecropin A Cecropin P1 Defensin NP-2 Lactoferricin B Lactoferrin Lysozyme Magainin 1 Magainin 2 Melittin Miotoxinin II Poly-L-lysine Polymyxin B Poly-L-ornithine EDTA Tris Lysosomal extract

ornithine lipids could also shield the negatively charged groups of Kdo and lipid A (Outer Membrane Topology), which are the initial targets of these agents (Fig. 20). In many Gram-negative organisms, such negatively charged groups are bridged by divalent cations that play an essential role in outer-membrane stability. However, the resistance of Brucella to divalent cation chelators shows no significant structural role for this effect, and it can be postulated that their absence is compensated by the stronger hydrophobic anchorage of the LPS (Velasco et al., 2000).

Brucella

Salmonella

Metabolism This section summarizes what is known about the uptake of nutrients (Extracellular Enzymes and Uptake), intermediary metabolism (Intermediary Metabolism), and energy-yielding processes (ATP Synthesis and Respiratory Chain) of Brucella and its response to environmental stress (Response to Environmental Stress). Recently, the results of a large genomic sequence survey carried out in B. abortus (http:// www.iib.unsam.edu.ar/genomelab/brucella/

362

E. Moreno and I. Moriyó n

index.html) (Sánchez et al., 2001) and the complete sequences of the genomes in B. melitensis strain 16M (http://www.genome.scranton.edu/ Brucella/) (DelVecchio et al., 2002) and B. suis (www.tigr.org/) (Paulsen et al., 2002) have been made available to researchers. As our knowledge of the Brucella genome expands, it is expected that many details of its physiology will be revealed and become accessible to experimental testing. Although reference to some of the sequence data will also be made, the reader has to keep in mind the problems associated with the interpretation of this still incomplete information. Extracellular Enzymes and Uptake The brucellae are devoid of extracellular hydrolytic enzymes acting on proteins, nucleic acids or polysaccharides. Accordingly, these bacteria depend exclusively on low molecular weight nutrients, such as sugars, amino acids or oligopeptides, etc., that are already present in the growth medium and that, depending on their hydrophobicity, cross the outer-membrane either through the porins or directly through the bilayer; both possibilities exist in Brucella (Permeability). The periplasmic space of many Gram-negative bacteria contains binding proteins that act coordinately with some cytoplasmic membrane transport systems. Attempts to isolate binding proteins from Brucella have been unsuccessful, which has been attributed to a tighter interaction of these proteins with the cell than those reported for other bacteria (Rest and Robertson, 1974), probably reflecting its outermembrane properties (Lipopolysaccharide and Resistance to Outer-membrane Destabilizing Agents). However, the Brucella genome contains genes whose products have characteristics suggestive of the periplasmic binding proteins involved in transport of oligopeptides, dipeptides and amino acids (Sánchez et al., 2001). In one case (protein P39), the genetic evidence has been complemented by modeling studies that also suggest a periplasmic binding role (De Fays et al., 1999) (Auxotrophic and Cell Cycle Genes During Intracellular Life). Sequences corresponding to aquaporins, mechanosensitive channels, transporters for the uptake of osmoprotectants, and probable ionspecific channels are also present. B. melitensis contains an ORF that is similar to the aquaporin of B. abortus (Rodriguez et al., 2000). There are genes coding for three putative ABC transporters for osmoprotectants, but not for members of the betaine/carnitine/choline transporter (DelVecchio et al., 2002). Uptake of Organic Nutrients The transport of glucose has been examined in B. abortus (Rest

CHAPTER 3.1.16

and Robertson, 1974) and the functional studies show the absence of a phosphotransferase (phosphoenolpyruvate-dependent) system and the presence of a proton-coupled uptake. The system is equally active in cells grown in the presence of fructose, galactose, erythritol and glucose, and seems therefore constitutive. Galactose acts as a competitive inhibitor for the uptake of glucose (Rest and Robertson, 1974) and complementation studies performed in E. coli with the B. abortus gene show that both sugars are transported by the same transmembrane protein (Essenberg et al., 1997). The putative glucosegalactose transport protein belongs to the proton-coupled transporters of the major facilitator superfamily, and a genomic sequence survey suggests ABC-type transporters active on sugars (D-fructose, D-mannose, melibiose, sucrose, trehalose, maltose, xylose, and sorbitol in B. melitensis), amino acids and oligopeptides (Sánchez et al., 2001; DelVecchio et al., 2002). The genomic analysis (B. melitensis) shows no erythritol-specific transporters (see below) such as those found in Rhizobium, but both glycerol and erythritol could be taken up by several putative polyol ABC transporters. A homologue to the gene coding for the multiple sugar-binding proteins of Agrobacterium and Azospirillum is present in at least B. melitensis. Also worth mentioning is that the bacA gene, which encodes a putative cytoplasmic membrane transport protein, is critical for virulence (LeVier et al., 2000). Uptake of Inorganic Nutrients Some aspects of the iron uptake systems of Brucella have been investigated. In early studies on the effect of iron availability, it was observed that Brucella releases large amounts of the monocatechol 2,3dihydroxybenzoate but not hydroxamate-type siderophores, and 2,3-dihydroxybenzoate and other simple monocatechols were observed to promote Fe uptake by an energy-dependent mechanism (Ló pez-Goñ i et al., 1992). Since monocatechols have much less affinity for iron than complex catechols (such as enterobactin or agrobactin) have, their role as siderophores is not obvious (for a possible different role of 2,3, dihydroxybenzoate, see Iron Chelation and Bactericidal Action). More recently, a B. abortus transposon mutant has been described that is unable to grow on iron restricted medium albeit it releases 2,3-dihydroxybenzoate (GonzálezCarreróet al., 2002). This mutant fails to release a minor catechol which, in cross-feeding experiments, has the ability to promote growth of the mutant under Fe limiting conditions. The mutated gene shows homology with genes (entC) involved in the synthesis of complex catechols by other bacteria, including vibriobactin and enter-

CHAPTER 3.1.16

obactin (González-Carreróet al., 2002). However, enterobactin fails to promote growth of B. abortus and, interestingly, neither agrobactin nor rhizobactin mediate Fe uptake by B. abortus (Ló pez-Goñ i et al., 1992). These data show that B. abortus produces a complex catechol siderophore (brucebactin) of unknown structure which is different from some of the catechols produced by related bacteria. A puzzling observation is that the release of catechols is not accompanied by the expression of new outer-membrane proteins that could act as siderophore receptors (Ló pez-Goñ i et al., 1992). However, because 2,3, dihydroxybenzoate and related compounds are moderately hydrophobic, direct penetration of siderophore chelates by the hydrophobic pathway could explain this observation. Once in the cytoplasmic compartment, iron has to be stored in such a way that the generation of free oxygen radicals, including the very toxic hydroxyl radical, is prevented, and a bacterioferritin gene with a high similarity to the corresponding E. coli gene has been characterized in B. melitensis (Denoel et al., 1995) (Iron Chelation and Bactericidal Action). Recently, some aspects of the high affinity system of nickel (Ni) uptake have been elucidated in B. suis (Jubier-Maurin et al., 2001). The nik gene cluster of B. suis closely resembles that of the corresponding E. coli operon, and although the regulatory gene (nikR) is found in a different position and orientation, both systems can complement the heterologous deficiency. This system is activated by low oxygen tension and metal ion deficiency and repressed when Ni is in excess. As expected (the enzyme contains Ni), its impairment leads to a severe reduction in urease activity (The Urea Cycle and Urease Activity). Finally, sequences compatible with systems involved in Mg++, Zn++, NO3− and SO42− transport have been identified in genomic surveys. Efflux Systems Genomic surveys (DelVecchio et al., 2002) show the presence of genes putatively coding for members of the cation diffusion facilitator (CDF) family and P-type ATPases involved in heavy metal efflux. Moreover, there are homologues of ABC transporters involved in drug efflux and of multidrug-resistance proteins and acriflavine-resistance and quaternary ammonium resistance systems. A TolC homologue (a common outer membrane component of several of these efflux pumps) is also present. Intermediary Metabolism Sugar Catabolism and Tricarboxylic Acid Cycle At least B. melitensis (strain 16M) carries genes predicted to code for the enzymes of all path-

The Genus Brucella

363

ways of central carbohydrate metabolism but those of the glycogen cycle. This includes genes for the pentose phosphate enzymes as well as for the complete Entner-Doudoroff pathway but this is in apparent conflict with the results of functional studies (DelVecchio et al., 2002). The pathways of glucose catabolism have been studied in B. abortus (strains 19 and 2308), B. melitensis (strain 16M) and B. suis (strain 1330) using glucose radiolabeled at positions C1, C2, C3, C4 and C6 in a complex medium (Robertson and McCullough, 1968a). The kinetics of 14CO2 release in dependence of the labeled position is C1 > C2 > C3 > C4 > C6, and this is compatible with either the operation of the hexose monophosphate pathway by itself or the hexose monophosphate plus the Entner-Doudoroff pathways working simultaneously, in both cases in conjunction with pyruvate oxidation through the tricarboxylic acid cycle. However, analysis of pyruvate from arsenite-poisoned cells or of alanine as the major product derived from pyruvate shows that glucose C1, C2 or C3 does not contribute significantly to the labeling of these two molecules. This implies that a C3–C4 cleavage does not occur significantly in the degradation of glucose and therefore that neither the EntnerDoudoroff pathway nor the hexose diphosphate pathway is active in Brucella. Differences between the 14CO2 release pattern of B. suis and the other species tested have been noted (Robertson and McCullough, 1968a) and attributed to a higher activity of the reactions leading from glyceraldehyde-3-phosphate to pyruvate. In cell-free studies with B. abortus 19, it has been observed that fructose 1,6-diphosphate is oxidized very slowly, that fructose diphosphate aldolase activity is very low and that, although phospho-2keto-3-deoxygluconate aldolase activity is detected, there is no phosphogluconate dehydratase activity (Robertson and McCullough, 1968b). Consistent with the genomic studies, the cell-free studies also show the major enzymatic activities of the pentose cycle when they are tested during anaerobic dissimilation of ribose5-phosphate. All these findings support the existence of an operative pathway in which glucose is phosphorylated (the glucokinase gene has been characterized; Essenberg, 1995) and oxidized first to gluconate-6-phosphate. Gluconate is then oxidized in the pentose cycle to yield 3 moles of CO2 plus 1 mole of glyceraldehyde-3-phosphate, and the latter is channeled into the tricarboxylic acid cycle via pyruvate (Fig. 25). So far, the oxidative pentose phosphate cycle as the major glucose catabolic pathway has been described in only a few bacteria. Glycerol is efficiently used by brucellae, and the genes that encode glycerol kinase and glycerol-3 phosphate dehydrogenase have been

364

E. Moreno and I. Moriyó n

CHAPTER 3.1.16 3 NAD+

ADP

ATP Glucose

3 Glucose-6-P

3 NADH2+

1

Erythritol

2

5 ATP

3 Gluconate-6-P

2 Fructose-6-P

ADP 2 NAD+ (Fp)

6 3

4

3 NAD+

2 NADH2+ (FpH2+) 2-Ribulose-5-P

3 NADH2+

CO2 1 Glyceraldehyde-3-P

Fig. 25. Glucose and erythritol catabolism in B. abortus. 1) glucokinase; 2) glucose-6-phosphate dehydrogenase; 3) gluconate-6phosphate dehydrogenase; 4) transaldolase and transketolase interconversions; 5) glucosephosphate isomerase; 6) erythritol pathway (see Fig. 26 and 27); and 7) terminal reactions of the glycolytic pathway and pyruvate dehydrogenase leading to the tricarboxylic acid (TCA) cycle.

3 CO2

7 TCA

identified in, at least, B. melitensis (DelVecchio et al., 2002). Also, early studies on the ability to grow on McCullough and Dick minimal medium (Table 9) with sucrose, glucose, galactose, mannose, fructose, threalose, xylose, arabinose and erythritol as the sole energy and carbon sources showed that all these compounds supported the growth of the four B. suis strains tested, that the growth of four B. melitensis strains showed variable results in arabinose, mannose, sucrose and threalose, and that none of the four B. abortus strains grew on arabinose, mannose, sucrose and threalose (McCullough and Beal, 1951). Because McCullough and Dick medium does not furnish the minimal requirements of the less prototrophic Brucella species (Nutritional Requirements), absence of growth in these experiments has to be interpreted with care. Nevertheless, the results suggest that at least B. suis has both transport and enzymatic abilities to channel all those compounds into the tricarboxylic acid cycle, and several putative sugar kinases can be identified in genomic surveys (Sánchez et al., 2001; DelVecchio et al., 2002). Similar speculations can be made on the meaning of the oxidative metabolic patterns established for the classical species and biotypes of Brucella (Table 2), taking always into account that the differences may also reflect differences in uptake and not necessarily in catabolic abilities. The tricarboxylic acid cycle has been demonstrated in B. abortus 19 by using pyruvate- and acetate-14C (Robertson and McCullough, 1968a) and also in other B. abortus strains in enzymatic studies (Altenbern and Housewright, 1953; Marr et al., 1953). Evidence for the functioning of several steps of the cycle has also been provided by studies on the oxidative dissimilation of amino acids by B. abortus. These data are consistent with those of the genomic analyses which also

show genes putatively coding for the enzymes required for anaplerotic reactions (NADdependent malate dehydrogenase [malic enzyme], phosphoenolpyruvate carboxykinase and pyruvate carboxylase). In addition, there is a pyruvate phosphate dikinase, which converts pyruvate to phosphoenolpyruvate (Oxidation of Amino Acids). Erythritol The early studies described in the section Sugar Catabolism and Tricarboxylic Acid Cycle showed that erythritol is able to support the growth of all B. abortus, B. suis and B. melitensis strains (McCullough and Beal, 1951) and this has been confirmed repeatedly. Other characteristics also make the metabolism of this compound by Brucella particularly interesting: growth with erythritol occurs earlier than with other substrates, and at least in B. abortus, erythritol is used preferentially over glucose in complex media (Anderson and Smith, 1965). These observations, and the presence of erythritol in the placenta of ungulates, led to the controversy on the possible relationship(s) between erythritol metabolism and virulence and viscerotropism in Brucella. The metabolic pathway of erythritol was elucidated in B. abortus B19 (British strain 19) (Sperry and Robertson, 1975). Meso-erythritol is oxidized through five steps that involve a kinase, three dehydrogenases and a decarboxylase to CO2 and dihydroxyacetone-phosphate, which is then converted to pyruvic acid through the final steps of glycolysis (Figs. 25 and 26). Two dehydrogenases are membrane bound and require NAD, but the third (3-keto-L-erythrose-4phosphate dehydrogenase) is coupled directly to the electron transport chain. The pathway is different from that described in other bacteria that either transform erythritol in L-erythrulose

CHAPTER 3.1.16 CH2OH

The Genus Brucella

ATP

ADP

1 (eriA)

HC-OH CH2OH

2 (eriB)

CH2OH

FpH2+

HO

C

HO

CH C

3-keto-L-erythronate-4-P

HO 3 (eriC)

CH2OH D-erythrulose-1-P

CH2OH

CH2O-P

HC

O

CH C

O

CH2O-P 3-keto-L-erythrose-4-P

CO2

O

O

O

HC-OH

D-erythritol-1-P

meso-erythritol

4 (eriD)

C

HC-OH

NAD+ NADH2+

CH2O-P

HC-OH

HC-OH

Fp

NAD+ NADH2+

CH2O-P

365

HC

O TCA

HC-OH

HC=O 5

6 CH2O-P dihydroxyacetone-P

7 CH2O-P glyceraldehyde-3-P

Fig. 26. The erythritol pathway in B. abortus. 1) erythritol kinase; 2) erythritol-1-phosphate dehydrogenase; 3) erythrulose1-phosphate dehydrogenase; 4) 3-keto-erythrose-4-phosphate dehydrogenase; 5) 3-keto-erythronate-4-phosphate decarboxylase; 6) phosphotriose isomerase; and 7) terminal reactions of the glycolytic pathway and pyruvate dehydrogenase leading to the tricarboxylic acid (TCA) cycle. Putative genes are indicated in parenthesis.

by means of a dehydrogenase (Acetobacter suboxydans and Alcaligenes faecalis) or split L-erythrulose-1-phosphate directly into formaldehyde plus dihydroxyacetone-phosphate (Propionibacterium pentosaceum). However, it has to be noted that the ability to use erythritol is also present in many bacteria of the α-2 subgroup of the Proteobacteria including strains of Rhizobium tropicii, Rhizobium loti, Sinorhizobium meliloti, Sinorhizobium fredii, Agrobacterium biovar 2 and all known species of the genus Ochrobactrum (De Lajudie et al., 1994; Lebhun et al., 2000). None of these bacteria are primarily associated with animals, and although the erythritol pathway has not been elucidated in them, it should not be different from that of Brucella. Therefore, it seems likely that the erythritol pathway of Brucella has not resulted from the adaptation to pathogenicity in ruminants, but represents an ancestral metabolic pathway for the α-2 Proteobacteria (Genome Evolution). In the study of over 100 strains of B. abortus, it was noted that erythritol not only did not promote growth of strain US19, but actually inhibited it (Jones et al., 1965). The US19, the current B. abortus 19 vaccine, and the B19, the British 19, are possibly related but whereas the former is unable to grow on erythritol which is, in fact, toxic for it, the latter catabolizes erythritol (Identification of Vaccine Strains). This peculiarity of the vaccine was explained in the course of the studies on erythritol metabolism. It was found that B. abortus US19 lacks D-erythrulose1-phosphate dehydrogenase activity and has a 20% reduced 3-keto-erythrose-4-phosphate

dehydrogenase activity (Sperry and Robertson, 1975). Presumably the build-up of intermediary metabolites becomes toxic for strain 19 and prevents growth when erythritol is present. This defect can be traced to a 702-bp deletion in the ery region of strain 19 (Sangari et al., 1994b) (Identification and Typing). This region is known to contain at least four genes of which presumably eryA codes for the erythritol kinase, eryB for the erythritol-1-phosphate dehydrogenase, eryC for the erythrulose-1-phosphate dehydrogenase and eryD for a regulator of ery expression (Sangari et al., 2000). The 702-bp deletion in strain 19 affects both eryC and eryD, thus creating the described phenotype. In contrast, B19 carries no deletion (Sangari, 1993), and “revertants” of US19 for which erythritol is not toxic probably carry compensatory mutations elsewhere in the chromosome. Erythritol analogues have been studied for their in vitro and in vivo anti-Brucella activity. Fluoro-mesyl and bromo-derivatives of erythritol and threitol inhibit the growth of B. abortus, B. melitensis and B. suis in vitro and in bovine phagocytes (typical concentrations causing a 50% inhibition of B. abortus range from 5 µg/ml [for 1,2-3,4-dianhydro-DL-threitol] to 100 µg/ml [for 1,4-dimesylerythritol]). The therapeutic value of the analogues in B. abortus-infected guinea pigs is, however, very limited (Smith et al., 1965). It has been shown that primers taken from the ery region amplify DNA of the expected size from B. ovis and B. canis (Sangari, 1993). These species do not or poorly oxidize erythritol (Table

366

E. Moreno and I. Moriyó n

CHAPTER 3.1.16

2), but this substrate is not toxic for them. Thus, their characteristics are different from those of B. abortus US19. There is a need for comparative studies to understand these differences. Oxidation of Amino Acids In a study of the ability of B. abortus 19 to grow on amino acids and related compounds, it was found that only glutamate, asparagine and histidine were used as the sole nitrogen source (Gerhardt et al., 1950b), and that the rate of oxygen uptake with glutamate was considerably greater than with any of the other substrates (Gerhardt et al., 1950a). It was also found that a simple medium containing glutamate as the nitrogen, carbon and energy source, plus accessory factors, supported growth (Table 9). The data obtained in subsequent metabolic studies (Marr et al., 1953) are consistent with a very active pathway through which glutamate is converted first to ketoglutarate and then metabolized in the tricarboxylic acid cycle to yield energy and reduced coenzymes plus intermediary metabolites for biosynthesis, including pyruvate. Oxidation of glutamate was also shown to be coupled to the urea cycle (The Urea Cycle and Urease Activity). Interest in the metabolism of glutamate was spurred by the observation that, at least in B. abortus and B. suis, its oxidative rate was lower in strains of high virulence than in strains of low virulence (Wilson and Dasinger, 1960). However, cell free studies showed no differences in the pathway of glutamate oxidation or in enzymatic activity associated with virulence, and it was concluded that unequal rates of permeability (Properties of the Outer Membrane) to the substrate could account for the observations (Dasinger and Wilson, 1962). The Urea Cycle and Urease Activity Brucella abortus and B. melitensis resting cells slowly metabolize glutamate with transient accumulation of arginine, ornithine and citrulline as metabolites, and at least in B. abortus, aspartate

ornithine lipids

+ ATP

6

Urea

Physiology of CO2 Requirements The biochemical basis of the CO2-dependence of some Brucella species and biotypes is not known. CO2 cannot be replaced by cell extracts, complex nutrients, tricarboxylic acid cycle intermediates, purines or pyrimidines, and the major products of CO2 assimilation in complex media are in the protein fraction as alanine and glycine and in the nucleic acid fraction as pyrimidines (Newton et al., 1954b; Newton and Wilson, 1954a). In keep-

NH4+ + CO2 + H2O

NH4+ + CO2 5

is also metabolized to ornithine (Cameron et al., 1952). Moreover, arginase activity (direct conversion of arginine into ornithine) exists in B. abortus, B. melitensis and B. suis (Cameron et al., 1952; Cameron et al., 1954). This enzymatic activity is considered the landmark of the urea cycle, and therefore, these results strongly suggest that at least these three Brucella species should have this metabolic cycle. The linkage with the metabolism of glutamate shown by the preceding observations could occur through the tricarboxylic acid cycle plus an aspartic-glutamic transaminase, since this enzymatic activity has been shown in at least B. abortus (Altenbern and Housewright, 1951). Moreover, Brucella contains ornithine lipids (Figs. 13 and 14) in the cell envelope and the cycle should furnish this amino acid for acylation and further modification. All classical Brucella smooth species, B. canis and some B. ovis strains (Corbel and Hendry, 1985) show a characteristic urease activity as do the Brucella marine isolates, and the enzyme is located in the cytoplasmic fraction (Brun and Descous, 1984). It has been proposed that urease is available in Brucella because of the occurrence of the urea as an intracellular metabolite (Cameron et al., 1952). Removal of urea would be necessary when the bacteria are growing with amino acids as the main source of carbon and energy. The role of urease in virulence is discussed elsewhere in the chapter (Metabolism), and these aspects of the metabolism of Brucella are summarized in Fig. 27.

Carbamoyl-P ornithine

H2O

2

1

Citrulline

Ariginine 4 Fumarate

Pi

3 Arginino-succinate AMP + PPi

Glutamate

Aspartate + 8 ATP

α -Keto-glutarate 7

CO2

Oxaloacetate

Fig. 27. The urea cycle and associated reactions in B. abortus. 1) arginase; 2) ornithine transcarbamoylase; 3) argininosuccinate synthetase; 4) argininosuccinase; 5) urease; 6) synthesis of membrane ornithine lipids; 7) tricarboxylic acid cycle reactions; and 8) glutamate-aspartate transaminase.

CHAPTER 3.1.16

ing with the strong respiratory activity of the Brucella (Physicochemical Requirements and Atmosphere), these observations demonstrate that CO2 is not required to reduce oxygen tension, but rather that it is needed as a nutrient per se. In bacteria, CO2 is used in the synthesis of purines, pyrimidines, carbamoyl-phosphate and fatty acids. In some cases studied (Burns et al., 1995), the CO2 requirement is due at least in part to the high Km of one of the carboxylases involved. The rate of appearance of CO2independent phenotypes in the CO2-dependent strains has been estimated in approximately 3 × 10−10 per cell division (Marr and Wilson, 1950). Therefore, if this Brucella phenotype is dependent on the affinity of carboxylases for CO2, few of them are involved. This is consistent with the observation that a CO2-dependent strain and its CO2-independent derivative did not differ in the distribution of fixed 14CO2 and showed the same basic nutritional requirements (Newton et al., 1954a; Newton and Wilson, 1954b). ATP Synthesis and Respiratory Chain Because glucose is mostly oxidized by the hexose monophosphate pathway, the substrate-level phosphorylation contribution to ATP synthesis should be minimal (provided glyceraldehyde 3phosphate is only an intermediary to the tricarboxylic acid cycle), and Brucella should obtain ATP from glucose or erythritol (and presumably other sugars) mostly by coupling coenzyme oxidation to the electron transport system (Rest and Robertson, 1975). In functional studies performed with the envelope fraction of B. abortus 19, NADH2+ dehydrogenase activity and also other primary dehydrogenases using L-lactate and succinate as substrates were observed by spectral analyses and polarographic determination of oxygen consumption. NADH2+ dehydrogenase and succinate dehydrogenase have also been demonstrated in the characterization of the cell envelope fraction of other B. abortus strains and in B. melitensis (Moriyó n and Berman, 1982). Moreover, at least in B. abortus, oxidation of erythritol 1-phosphate is also directly coupled to the electron transport chain (Rest and Robertson, 1975) through the 3-keto-l-erythrose 4-phosphate dehydrogenase (Sperry and Robertson, 1975; Fig. 26). Analysis of the free lipid fraction of B. melitensis and B. abortus (Thiele and Schwinn, 1969; 1973) shows the characteristic e− /H+ transporter ubiquinone Q10, and the polarographic data obtained with B. abortus 19 (Rest and Robertson, 1975) are consistent with the location of the primary dehydrogenases early in the respiratory chain. The spectrophotometrical analyses also provided evidence for the presence of flavoproteins and several cytochromes which were different in exponentially growing bacteria

The Genus Brucella

367

from those in the late exponential phase of growth, suggesting the ability to adapt the terminal elements of the electron transfer to different oxygen tensions (Rest and Robertson, 1975). According to the predictions of the gemomic analysis, in B. melitensis a terminal cytochrome o ubiquinol oxidase acts under normal oxygen concentration and a cytochrome cbb3 oxidase acts under low oxygen concentration. During microaerobic conditions, B. melitensis can use the cytochrome bd quinol oxidase for electron transfer to oxygen from the membrane quinone pool. Complete sets of genes for all of the subunits for complex I (NADH-quinone oxidoreductase), complex II (succinate dehydrogenase), and complex III (ubiquinol-cytochrome c reductase) and for the soluble electrontransporting cytochromes c2 and c552 have also been identified as well as those of several primary dehydrogenases (D- and L-lactate dehydrogenases, flavoprotein-quinone oxidoreductase, glycerol-3-phosphate dehydroand malate:quinone genase [NAD(P)+], oxidoreductase). These enzymes supply electrons and protons for the aerobic respiratory chain and oxidative phosphorylation pathways. Genes coding for H+-coupled ATP synthase, NAD(P) trans-hydrogenase, and Na+/H+ and K+/H+ antiporters are present and the cognate proteins should be involved in oxidative phosphorylation and the proton gradient maintenance. Nitrate-nitrite reduction is observed in all Brucella classical species but B. ovis. Nitrate reductase activity was detected in the course of the above-described studies on the characterization of the electron transport system of B. abortus 19 (Rest and Robertson, 1975). Moreover, in the studies on the catabolism of erythritol, it was shown that nitrate was almost as effective as oxygen in stimulating the breakdown of erythritol, whereas nitrite and hydroxylamine had no effect (Sperry and Robertson, 1975). Apparently, B. abortus (and possibly the rest of the Brucella species reducing nitrate to nitrite in a dissimilative way) has a terminal nitrate reductase coupled to the electron transport chain via flavoproteins and cytochromes and this is consistent with the genomic analyses. At least in B. melitensis, the respiratory nitrate reductase complex consists of four subunits encoded by a single operon (DelVecchio et al., 2002). Response to Environmental Stress Aerobic organisms are endowed with enzymes that detoxify the reactive by-products of oxygen, which are generated mostly during the operation of the respiratory chain. Two types of superoxide dismutase (Sod) have been found in Brucella abortus, a cytosolic Mn++-Sod and a Cu++/Zn++-Sod

368

E. Moreno and I. Moriyó n

(Bricker et al., 1990) located in the periplasmic space (Stabel et al., 1994). By Western-blotting with antisera to the homologous E. coli protein, the periplasmic Sod has been demonstrated in all Brucella species and biovars tested except in B. neotomae and B. suis biovar 2 (Bricker et al., 1990). Moreover, catalase activity can be demonstrated in all Brucella species in the standard H2O2 assay, and according to manometric studies performed with B. abortus, B. melitensis and B. suis, the latter species displays the strongest catalase activity, which is also slightly higher in B. melitensis than in B. abortus (Huddleson and Stahl, 1943). A periplasmic catalase has been purified and cloned from Brucella abortus and characterized as a tetrameric protein (MW of the monomer 55,000) lacking a typical N-terminal export signal sequence (Sha et al., 1994). The periplasmic location of catalase and Cu++/ Zn++-Sod suggests that they protect the cell from external sources of oxidative damage (Stabel et al., 1994), such as those generated by phagocytes upon ingestion of bacteria (Neutralization of Oxygen and Nitrogen Intermediates). In fact, the periplasmic catalase behaves as an oxidative stress protein, and exposure of B. abortus cells to sublethal concentrations of H2O2 increases both their resistance and the expression of the enzyme. Although the magnitude of the increase is lower, oxidative stress also increases expression of the periplasmic Cu++/Zn++-Sod (Table 15). Because Sod activity leads to the production of H2O2, this different level of response is consistent with the respective roles of these enzymes in protecting the cell from oxidative damage (Kim et al., 2000). Homologues to the htrA, groEL and dnaK genes have been identified in Brucella, and the latter has been cloned and shown to express a functionally active product in a thermosensitive E. coli dnaK mutant (Cellier et al., 1992; Elzer et al., 1994; Lin et al., 1992; Roop et al., 1994; Tatum et al, 1994). As expected, the synthesis of the two cytoplasmic molecular chaperones (GroEL and DnaK) and the periplasmic protease HtrA is stimulated under heat stress (42°C), and impaired growth has been shown for an htrA-defective mutant under these conditions. Moreover, an increased expression of these three genes is also observed in acidified media (Table 15). In addition to the above-described proteins, heat, oxidative, acid and nutritional stresses cause an increase in the synthesis of over 100 proteins, among which many are not expressed under normal conditions. Conversely, many proteins expressed in normal media become repressed under these conditions. A small fraction of these proteins has been identified by proteomic methods (Table 15), and some elicit

CHAPTER 3.1.16

antibodies in naturally infected animals, demonstrative of their expression in the host. It is noteworthy that the profile of proteins induced and repressed when B. abortus grows in cultured macrophages does not correspond to a simple sum of the proteins observed under stress conditions in vitro (Lin and Ficht, 1995; Rafie-Kolpin et al., 1996).

Genetics The Brucella Genome The genome of a bacterium includes the chromosome plus any extrachromosomal DNA, such as prophages and plasmids carrying nonessential genes useful only under some environmental conditions. Bacterial chromosomes can be linear and bacteria can have more than just a single chromosome (Kolst, 1999). Interestingly, of the bacteria currently known to have a large part of their essential genes in replicons outside the larger chromosome (i.e., to have one or more additional chromosomes of different size), many belong, like Brucella, to the α Proteobacteria (Jumas-Bilak, 1998b; Kolst, 1999; Moreno, 1998). Moreover, some members of the α Proteobacteria are known to have a linear chromosome in addition to the major circular one (Jumas-Bilak et al., 1998b) and many also have plasmids (Moreno, 1998). In this taxonomic context, the brucellae are characterized by the presence of two or, in B. suis biovar 3, one circular chromosome and by the absence of plasmids (Fig. 1). The results of B. melitensis genomic sequence (DelVecchio et al., 2002), a survey carried out in B. abortus (Sánchez et al., 2001) and the genome sequence of B. suis (Paulsen et al., 2002) represent a wealth of genetic details yet to be evaluated. The general genetic characteristics of the Brucella genome are summarized here, and their evolutive and taxonomical meaning is discussed elsewhere in the chapter (Biosafety).

Brucella Chromosomes Number and Species Differences The distribution of some essential genes (ribosomal rrn and genes coding for heat shock proteins), combined with pulse-field gel electrophoresis, shows that the reference strains of all B. melitensis and B. abortus biovars, B. suis biovar 1, B. ovis, B. canis, and B. neotomae carry two circular chromosomes of about 2.1 and 1.15 Mb (MichauxCharachon et al., 1993; Jumas-Bilak, 1998a; Jumas-Bilak, 1998b), and the reference strains of B. suis biovar 2 and biovar 4 also have two circular chromosomes but of 1.85 and 1.15 Mb (Jumas-Bilak, 1998a). On the other hand, the reference strain of B. suis biovar 3 has a single

Repressed

Induced

Repressed

Induced

Periplasmic serine protease (DEGP_BRUAB) Amino acid-binding periplasmic protein (85.7%; Rhizobium leguminosarum; AAPJ_RHILV) Ribosome releasing factor (RRF_BRUME) Superoxide dismutase (73.1%; Salmonella; SODF_SALTY) Electron transfer flavoprotein α-subunit (80%; Bradirhizobium japonicum; ETFA_BRAJA) ATP-dependent Clp protease (heat-shock protein) (85.7%; E. coli; CLPP_ECOLI) Iron storage protein (BFR_BRUME) Regulator element of a two-component system involved in virulence (O67996) Homologous to an invasion-associated protein (60%; Bartonella bacilliformis; IALB_BARBA) (66.7%; Mycoplasma genitalium; ODPB_MYCGE) Periplasmic enzyme (CATA_BRUAB) Periplasmic enzyme (SODC_BRUAB)

HtrA

DnaK GroEL HtrA Malate dehydrogenase

30S ribosomal protein S1

Pyruvate dehydrogenase E1 component β-subunit Catalase Cu-Zn SOD

IalB

Bacterioferritin BvrR

ClpP

α-ETF

Fe and/or Mn SOD

RRF

mRNA binding protein (64.3%; Rhizobium meliloti; RS1_RHIME) Also increased by heat stress (see above) Also increased by heat stress (see above) Also increased by heat stress (see above) Krebs cycle enzyme (100%; Rhizobium leguminosarum; MDH_RHILV)

Molecular chaperone (CH60_BRUAB)

GroEL

AapJ

Molecular chaperone (DNAK_BRUOV)

Characteristicsa

DnaK

Protein

+++ +++ ++ +

++

Denoel et al., 1997 Teixeira-Gomes et al., 1997, 2000

+

Teixeira-Gomes et al., 1997, 2000 Rafie-Kolpin et al., 1996 Rafie-Kolpin et al., 1996 Teixeira-Gomes et al., 1997; 2000

Kim et al., 2000 Teixeira-Gomes et al., 1997, 2000; Tabatabai and Hennager, 1994; Kim et al., 2000 Teixeira-Gomes et al., 1997, 2000

Teixeira-Gomes et al., 1997, 2000

Teixeira-Gomes et al., 1997, 2000

Teixeira-Gomes et al., 1997, 2000

Teixeira-Gomes et al., 1997, 2000

Vizcaino et al., 1996; Teixeira-Gomes et al., 1997, 2000 Teixeira-Gomes et al., 1997, 2000

Teixeira-Gomes et al., 1997, 2000; Cellier et al., 1992 Lin et al., 1992; Kittelberger, 1995a; Lin and Ficht 1996; Teixeira-Gomes et al., 1997, 2000 Roop et al., 1994; Elzer et al., 1994; RafieKolpin et al., 1996 Teixeira-Gomes et al., 1997, 2000

References

+

+

++

++

+++

+++

Immunoresponse in natural hosts

The Genus Brucella

Symbols: +, mild, ++, moderate, and +++, strong immunoresponse(s). a Where appropriate the % identity, bacteria, and/or ID in SWALL protein database is included in parentheses.

pH 5.5

Oxidative

Induced

Heat

Repressed

Regulation

Stress

Table 15. Some proteins of Brucella that are regulated by heat, oxidative and pH stress.

CHAPTER 3.1.16 369

370

E. Moreno and I. Moriyó n

circular chromosome of 3.10 Mb (Jumas-Bilak, 1998a; Fig. 1). Brucella marine strains also possess two chromosomes close to the size of B. abortus (D. O’Callaghan, personal communication). For the species carrying the 2.1- plus 1.15-Mb chromosomes, the physical maps demonstrate a high conservation of restriction sites, although with differences due to small insertions and deletions of 1 to 32 Kb (Michaux-Charachon et al., 1997), accounting for the species-specific restriction patterns (Table 4). On the other hand, there are major chromosomal rearrangements affecting the 1.85- and 1.15-Mb chromosomes of B. suis biovar 2 and biovar 4, since sequences of the 2.1 Mb chromosome of B. suis biovar 1 are found in the 1.15 Mb chromosome of these biotypes and also in the single chromosome of B. suis biovar 3. Similar to what occurs in other bacteria, these differences in chromosome size and number can be explained by rearrangements caused by homologous recombination at the redundant rrn loci (Jumas-Bilak, 1998a) or insertion sequences (reviewed by Moreno, 1998). Taking into account the great homogeneity of the genus Brucella (Ancestor-descendent Relationships), the variations observed in B. suis are remarkable. For hypotheses on the origin of the two chromosomes, see Genome Evolution. Size and Percentage of Guanine Plus Cytosine The data summarized in Table 1 (Number and Species Differences) show that the size of the genome varies between the 3.25 Mb of most Brucella spp. and the 3.0 Mb of B. suis biovar 3. This is significantly less than the 5.29 Mb (2.70 and 1.90 Mb chromosomes, plus two 0.59- and 0.10-Mb plasmids) of the genome of the closest known relative of Brucella, the genus Ochrobactrum, and the differences are due to both the smaller size of the chromosomes and the absence of plasmids in Brucella. Similar or even larger differences in genome size exist with other taxonomic neighbors, such as Agrobacterium, Rhizobium, Phyllobacterium and Bradyrhizobium, and the average G+C content of Brucella spp. (57%) is somewhat lower than that of these bacteria (Table 1). On the other hand, a relatively reduced genome size and a comparatively low G+C content are also characteristic of other pathogenic α Proteobacteria (Fig. 1) and probably relate to their adaptation to an intracellular or pericellular animal environment (Genome Evolution). Methylation and Control of Chromosome Duplication and Cell Cycle It is not known how segregation of the genome is controlled in Brucella or in other bacteria carrying more than one chromosome. However, it is known that

CHAPTER 3.1.16

Brucella has genes whose products are involved in the control of the cell cycle. In bacteria, DNA methylation not only is involved in restrictionmodification systems but also plays regulatory roles in the cell. The Dam methyl transferases of the γ Proteobacteria control the timing of initiation of chromosome duplication and the expression of some genes, including virulence genes, and are involved in methyl-directed DNA mismatch repair (Reisenauer et al., 1999). The counterpart of Dam in the α Proteobacteria seems to be CcrM (cell-cycle-regulated methyl transferase). This enzyme was originally described in Caulobacter crescentus, a bacterium that presents a morphologically defined cell cycle Caulobacter and of which synchronic cultures are relatively easy to obtain. The CcrM catalyzes the methylation of the adenine residue in GANTC sequences, and like Dam, uses Sadenosylmethionine as a donor and has no known cognate restriction enzymes. CcrM (but not Dam) is essential for cell viability and its activity is controlled during the cell cycle by tightly regulating the activation of ccrM expression (by the global regulator CtrA [see below] and quick constitutive degradation by Lon proteases). When ccrM is transcribed constitutively and the chromosomes are fully methylated, most Caulobacter cells abnormally increase the number of chromosomes from 1 or 2 to 3, and methylation seems to be particularly frequent in close proximity to DnaA boxes. On these bases, it has been postulated that CcrM plays a key role in the regulation of the chromosome replication (Reisenauer et al., 1999). B. abortus carries a CcrM homologue (78% similarity) that has been shown to be essential for viability, and when the number of ccrM copies is artificially increased over a given number, a dramatic effect in cell shape and a relaxation in the control of DNA are observed; cells appear highly branched and contain up to five genome equivalents (Robertson et al., 2000b). Moreover, B. abortus constructs overexpressing ccrM to levels not inducing aberrant morphologies or altered chromosome number still present reduced rates of intracellular multiplication (Robertson et al., 2000b). This indicates that other cell functions related to intracellular survival are likely to be regulated by CcrM. The attenuation of virulence caused by these alterations in CcrM is discussed elsewhere (Auxotrophic and Cell Cycle Genes During Intracellular Life). Insertion Sequences (IS) and Other Genetic Elements All classical Brucella spp. examined carry a characteristic insertion sequence (IS711 or IS6501) that is also present in the marine mammal Brucella isolates examined so far (Clavareau et al., 1998; Halling and Zehr, 1990;

CHAPTER 3.1.16

Regulation of Gene Expression The few examples of known regulatory systems of Brucella that have been partially characterized are briefly discussed here. The BvrR-BvrS System This is a twocomponent regulatory system. The sensor protein kinase BvrS is homologous to sensor proteins of the histidine protein kinase family, and the response regulator BvrR shows significant similarity to response regulators of the OmpR subfamily. The BvrS protein bears the

371

characteristics of an intrinsic cytoplasmic membrane protein with a periplasmic domain that, in all likelihood, carries the region sensing an unidentified environmental stimulus. This protein crossreacts with ExoS sensor protein of Rhizobium meliloti (C. Guzmán-Verri et al., unpublished observations). Thus, it is presumed that, under some environmental circumstances, BvrS catalyzes its own phosphorylation and then transfers the phosphoryl group to BvrR, which, in turn, coordinately activates or represses the expression of genes that, although not identified as of yet, have been demonstrated to be essential for virulence and intracellular multiplication (Sola-Landa et al., 1998; Mechanisms of Entry to Host Cells). Dysfunction of the BvrS-BvrR system is known to alter the outer-membrane permeability, the expression of several Omp3 proteins (among the most conspicuous Omp3a [Omp25] and Omp3b) (Outer Membrane Proteins), and the pattern of lipid A acylation (C. Guzmán-Verri et al., unpublished results), but the underlying genetic mechanisms are yet to be unraveled (Fig. 28; The Brucella Genome). An important feature of the BvrS-BvrR system is that it is highly conserved in other pericellular pathogens or endosymbionts of the α-2 Proteobacteria; it is homologous to the ChvI-

Omp3B

Core

Omp3A

Outer membrane homeostasis

External signal

Lipid A

S

P

his

P R

P

Outer membrane homeostasis

Halling et al., 1993; Ouahrani et al., 1993). In addition, there are other IS elements scattered in the genome of those brucellae that have been sequenced (DelVecchio et al., 2002; Paulsen et al., 2002). The insertion sequence IS711 is bound by two 20-bp imperfect inverted repeats, has a G+C content similar to that of the genome and is of 842–836 bp in length. It shows significant similarity (53.4%) with IS427 of A. tumefaciens, suggesting a common ancestral sequence, and like some other IS, has overlapping open reading frames rather than one long open-reading-frame extending over the length of the element. A long open-reading-frame of 708-bp encodes a putative protein sharing sequence identity with the hypothetical protein 1 of A. tumefaciens and with a transposase of Mycobacterium tuberculosis. Characterization of the flanking sequences suggests that some IS711 copies may have resulted from mechanisms other than transposition and also that there are hot spots for insertion. Two repeated palindromic DNA elements (Bru-RS1 and Bru-RS2) that are present in more than 35 copies in brucellae act as such insertion hot-spots (Halling and Bricker, 1994). The number of IS711 copies per genome varies among Brucella species: there are 20–35 copies in B. ovis and 5– 15 copies in other species. The copies present in a given strain may have minor differences in nucleotide sequence, and variations in position and number among species and biotypes result in genetic polymorphism (Table 4). At least in one case, the position of IS711 causes phenotypically demonstrated differences in protein expression between B. ovis and the other classical Brucella species (Halling and Zehr, 1990), suggesting that this insertion sequence is a significant source of natural genetic variation in Brucella (Genome Evolution). This hypothesis is also supported by the fact that the rough phenotype of B. abortus RB51 is due to the spontaneous insertion of IS711 in the wboA LPS gene (B- abortus RB51). The movement and transposition of this IS may be one of the sources for gradual microevolution, generating strains and eventually new species (Genome Evolution).

The Genus Brucella

Transcriptional activation or Transcriptional represion ? Fig. 28. Hypothetical model of the BvrS-BvrR regulatory system. Accordingly, an external unknown signal stimulates the sensor region of the BvrS protein, translocating the signal to a histidine kinase domain, which in turn promotes its own phosphorylation to phosphorylate the BvrR regulatory protein. The phosphorylated BvrR protein induces or represses transcription of genes not yet identified. By this system, the structure of the lipid A and the expression of Omp3a (Omp25) and Omp3b as well as the presence of other proteins at the Brucella outer membrane are regulated. The consequence of mutating either of these two genes in Brucella is an altered sensitivity to cationic peptides and hindrance to the cell invasion and intracellular replication.

372

E. Moreno and I. Moriyó n

ChvG (response regulator and sensor histidine protein kinase) of A. tumefaciens and to the ExoS of R. meliloti (Derivation of Ancestral Preexisting Structures for Virulence) as well as to genes found in Bartonella spp. The FeuP-FeuQ System A response regulator sequence with 96% similarity to R. leguminosarum FeuP has been identified in Brucella. In R. leguminosarum, the FeuP is part of a twocomponent sensory regulatory system (FeuPQ) operating as described for the BvrS-BvrR system and known to take part in the regulation of iron uptake. Its precise role in Brucella remains to be determined (Dorrell et al., 1998). The NtrBC System This is also a twocomponent sensory-regulatory system, with a sensor histidine-kinase protein (NtrB) and a regulatory transcriptional activator (NtrC) that has been identified in B. melitensis and B. suis (Dorrell et al., 1999). NtrC is a member of a family of bacterial proteins that enhance transcription of promoters recognized by the RNA polymerase carrying the alternative σ54 factor. The NtrCB system is involved in the regulation of the metabolism of nitrogen in several bacteria. When the NtrC element becomes growthlimiting, the genes necessary to use a variety of nitrogen sources become activated by NtrC. Consistent with this, B. suis NtrC mutants have increased metabolic activity in the presence of Lasparagine and L-alanine and decreased activity in the presence of other amino acids, such as glutamic, arginine, and lysine, and several sugars (Dorrell et al., 1999). The CtrA Regulator The CtrA regulator is a member of the response regulator family whose members, upon phosphorylation by a kinase, act as gene transcription activators. Brucella abortus carries a C. crescentus CtrA homologue (80% similarity) and the purified C. crescentus protein recognizes sequences upstream of the transcription starting site of the Brucella ccrM promoter, further stressing the similarity of the two methylating systems (Robertson et al., 2000b). By indirectly controlling the level of DNA methylation, CtrA should also be involved in the regulation of Brucella genes other than ccrM, but the kinase acting on CtrA has not been identified.

Genetic Exchange, Plasmids and Lysogenic Phages No naturally occurring system for genetic exchange has ever been demonstrated for any Brucella spp. Moreover, the weight of the evidence shows that, in contrast to their phyloge-

CHAPTER 3.1.16

netic neighbors, the brucellae do not harbor plasmids. They have not been found in an extensive search carried out with over 600 Brucella strains by a variety of methods (Meyer, 1990) and have never been occasionally observed. Likewise, no firm evidence has ever been provided on the existence of lysogenic phages (The Brucellaphages). Undoubtedly, these genetic characteristics relate to the peculiar intracellular habitat and to the genetic homogeneity of the group (Derivation of Ancestral Preexisting Structures for Virulence), and in all likelihood, the absence of plasmids is one factor contributing to the stability of the antibiotic sensitivity pattern of Brucella (Treatment). On the other hand, even with the limited information examined so far, there is evidence for gene acquisition by horizontal transfer. This is at least the case of the genes of the wbk region of B. melitensis 16M (the gmd, perA, wxm, wzt, wbkA, wbkB and wbkC LPS biosynthetic genes [Table 5]), the G+C content of which (44–49 mol%) significantly differs from that of the whole genome (57%) and suggests a shared origin with LPS genes of bacteria showing structurally related Opolysaccharides (Table 6; Acquisition of Ancestral Genes by Horizontal Transference).

Epidemiology Brucella is the causative agent of brucellosis, undulant fever, contagious abortion and Malta fever, all names that describe an important malady of animals and humans. When affecting domestic animals, primarily bovines, caprines, ovines, and swines, the disease acquires a great economical importance. If the bacterium is transmitted from animals to humans, then an important zoonosis is established and public health interest arises, mainly due to the fact that the disease causes disability and requires long and expensive treatment (Treatment). Since the time the bacterium “Micrococcus melitensis” (Brucella melitensis) was recognized more that 100 years ago in Malta as the causative agent of brucellosis, the epidemiology of this disease has received much attention. Many of the investigations carried out for already more than ten decades have been crucial for the control and eradication of brucellosis in several parts of the world. However, brucellosis remains as a relevant disease of domestic animals and as an important zoonosis, mainly in low-income countries (Geographical Distribution). Furthermore, brucellosis is an important infection of various wild life animals and a threat if some of the strains are utilized as biological weapons. In this respect, much controversy has come out in how to control and eradicate Brucella from these nat-

CHAPTER 3.1.16

The Genus Brucella

ural reservoirs and how to neutralize the bacterium if used in a confrontation (Brucella as a Biological Weapon). Because of these, the epidemiology and vaccination of brucellosis still capture the attention of many investigators in both developed and low-income countries (Vaccines and Vaccination). For extensive reviews in the epidemiology of animal brucellosis, the publications of Crawford and Hidalgo (1977) and Nielsen and Duncan (1990b) are recommended. For a comprehensive analysis on human brucellosis, the reviews of Spink (1956) and FloresCastro and Baer (1979) are recommended.

Geographical Distribution Although Brucella organisms infect a broad range of animals, the bacterial species and strains have strong tendency to remain in their pre-

373

ferred hosts (Biological Attributes in the Context of Brucella Species Definition). When infecting a secondary host (i.e., B. melitensis in cows), the removal of the source of infection (i.e., B. melitensis-infected goats) results in the disappearance of the bacteria from the secondary host population (Crawford and Hidalgo, 1977; Nielsen and Duncan, 1990b). In this respect, Brucella species and biovars follow the geographical distribution of their natural hosts and not the secondary or the accidental hosts, such as wild animals, humans or horses (Fig. 29). Since in many cases where domestic animals reside borders wildlife habitats, wild animals frequently become infected after contact with domestic livestock or their products, such as fetuses or infected tissues (Crespo-Leó n, 1994). In wild animal accidental hosts (i.e., coyotes), the disease is limited to the infected individual;

Distribution of caprines in the world

Caprines

Repotred cases of human brucellosis in Mediterranean countries in1996 FRANCE GREECE LEBANON TUNESIA Fig. 29. Geographic distribution of domestic caprines in the world, and human brucellosis in Mediterranean countries for the year 1996. Human and animal brucellosis caused by B. melitensis follows the distribution of goats, the natural host for this bacterium. From the United Nations Food and Agriculture Organization/World Health Organization/International Office of Epizootic, Animal Health Year Books.

PORTUGAL EGYPT JORDAN ALGERIA ITALY SPAIN SAUDI ARABIA SYRIA TURKEY 0

2000

4000

6000

8000

10000

374

E. Moreno and I. Moriyó n

however, when wild ungulates are involved (i.e., bison), the bacteria may remain for prolonged periods in the population. In this last case, the wild animals may be considered reservoirs of the disease (Davis, 1990; Hayes, 1977). Then, a double hazard occurs: one for wildlife and the other for the domestic animals. Consequently, a conflict between environmentalists and producers arises (McCorquodale and DiGiacomo, 1985). This pattern is more critical in developing countries, mainly in Africa and Asia, since there are not efficient procedures for the control and eradication of Brucella in wild animal populations. Bovine, ovine, caprine and swine brucellosis has also been eradicated from domestic livestock in Canada, Central Europe, Scandinavian countries, Japan and in the northern and central regions of the United States (Crawford et al., 1990; Crespo-Leó n, 1994; Plommet, 1992). In Australia and New Zealand, the prevalence of smooth Brucella pathogens in bovine, porcine, ovine and caprine herds is close to zero. Although at a low prevalence, B. ovis remains in ovine herds in some of those countries with extensive wool production (Crespo-Leó n, 1994). In Southeast Asia, bovine and caprine brucellosis is not common, although porcine brucellosis shows a relatively higher prevalence. Bovine, caprine and swine brucellosis also has been eradicated in Uruguay and in some regions of Argentina and Chile. However, ovine and caprine brucellosis caused by B. ovis and B. melitensis remains in some isolated herds in the latter two countries. In some of the “brucellosis-free” countries and regions, B. abortus biovar 1, B. canis, B. suis biovars 2 and 4 and B. neotomae prevail in wildlife hosts, mainly in bison, dogs, wild boars, reindeer and wood rats, respectively, animals native to these countries. In addition, marine Brucella strains infect seals, dolphins, porpoises and whales that live in the ocean limits and coasts of many of these “brucellosis-free” countries. In this sense, it could be said that with a few exceptions, Brucella pathogens are found worldwide (Jahans et al., 1997). Brucellosis caused by B. melitensis is endemic in Mexico and the principal cause of recurrent infections in caprine and ovine herds as well as the main zoonosis (Crespo-Leó n, 1994). In this North American country, B. melitensis is also a significant cause of bovine brucellosis due to the breeding of bovines close to goats and sheep. Brucella abortus, B. suis and B. canis are also prevalent as the main cause of brucellosis in bovines, swines and canines, respectively, in populations where goats are excluded. Brucella suis may be the second cause of occupational zoonosis in Mexico. In Central America, B. abortus (biovar 1) is the leading cause of brucellosis in bovines. Brucella melitensis is less frequently

CHAPTER 3.1.16

found in these animals because of the fewer goats and sheep found in these countries (Crespo-Leó n, 1994). Brucella abortus (biovar 1) is also the main cause of bovine brucellosis in most South American countries. Peru and Bolivia possess a large number of goats infected with B. melitensis, and the domestic livestock is the source of infection for American camelids (llamas and alpacas). Brucella suis has been sporadically reported in some areas of Brazil, Uruguay, Argentina and Central America; however, the prevalence is low because of the introduction of intensive management and shortterm recycling of swines in these countries. In the European Mediterranean countries, B. melitensis and B. ovis are more prevalent than other species of Brucella (Plommet, 1992). The northern regions of Portugal, Spain, and Italy, as well as Southern France, have strict control programs for eradication of B. abortus bovine brucellosis. In these countries, B. suis is detected sporadically and B. canis is not currently isolated and may have been eradicated. Among the Mediterranean countries in Europe, Turkey, Greece, Italy, Portugal and Spain have a higher incidence of brucellosis, which is largely caused by B. melitensis. In the Eastern European countries and countries of the former Soviet Union, mainly in the Balkanic areas, B. abortus and B. suis seem to be highly prevalent, although accurate numbers are not available. In some of these countries, in particular those located in the Southern Euroasiatic region in which caprine and ovine herds are maintained, B. melitensis remains an important cause of infection. Brucella suis biovar 4 is prevalent in reindeer and caribou living in the northern latitudes and are predominantly limited to Alaska, Canada and Siberia. Brucella melitensis is endemic in the Middle East, Arab countries, Indian Peninsula and southern Asia, mainly due to the population of caprines in these regions. Brucella abortus is also present in most of these countries and seems to be more prevalent in Central and Northern Asia (CrespoLeó n, 1994). In these countries, B. suis seems to be sporadic, possibly owing to the fact that the population of swines remains small because of religious practices. Brucella suis biovar 3 is prevalent in Southeast Asia, the Pacific Islands, Singapore, the Philippines and southern China, and Taiwan, whereas biovar 1 is more frequently found in Indonesia. Although it is known that the major pathogenic Brucella species are present in Africa, their prevalence and exact distribution are not known (FAO/WHO, 1986). The prevalence of B. melitensis in Africa seems to coincide with the distribution of the large number of caprines existing in many countries of this continent. Brucella abortus has been detected in bovine herds and in African wildlife, mainly in

CHAPTER 3.1.16

ungulates; however, other mammals, such as elephants, giraffes and camels, have been infected, mainly as a result of contact with domestic livestock. Brucella suis also has been detected in feral swines in several African countries. Although B. canis has been detected in widely separated areas, such as Mexico, Madagascar and Japan, the prevalence and exact distribution in the world are not known. It is likely that B. canis attains high prevalence in countries possessing a large population of dogs, mainly in developing countries where most dogs freely roam around urban and suburban areas (Carmichael, 1990).

Infecting the Host It is generally accepted that the most frequent route of infection is through the digestive tract. In goats and probably in other animals as well, the ingested bacteria are engulfed and transferred to the submucosa by intestinal M cells (Brucella Invades Healthy Hosts), but the ability to penetrate through the oral and pharyngeal mucosa has also been documented (Enright, 1990b). The conjunctival route seems to be an alternative path for natural infection in herds where intensive management of animals is a common practice (Plommet, 1977). The tissues of the aborted fetuses are massively contaminated with Brucella organisms (up to 1010 organisms per cm3 of tissue or allantoic fluid) and therefore are the main source of infection for susceptible hosts. Vaginal secretions are also an important source of infection, mainly for those animals under intensive management and when conditions of close contact among animals prevail. The udder of infected cows, goats and ewes frequently harbors a large number of Brucella organisms and the milk is a frequent source of infection. Although calves have been estimated to be more resistant to brucellosis than are adult animals (Russell, 1977), they can be infected by ingesting milk from infected cows, mostly after abortion or parturition. A similar situation may occur with kids, lambs and piglets. In ungulates, the infection is normally acquired through the habit of licking and smelling the aborted products and other infected animals. The congenital route is of epidemiological relevance because animals born from infected mothers may remain latently infected and without symptoms or serological response until pregnancy (Plommet, 1977; Fig. 9). Brucella organisms are also transmitted by the venereal route. This channel is more important in swine, ovine and canine brucellosis. Artificial insemination of contaminated semen with Brucella organisms is a risk factor for iatrogenic transmission (Manthei et al., 1950). Although it has been demonstrated that the bacteria can survive in water, soil and manure for

The Genus Brucella

375

limited periods of time and that transmission may occur by penetration of the bacteria through broken skin or by insect bites, these probably are more fortuitous than regular routes and are considered of minor epidemiological importance (Meyer, 1977). Rodents and carnivores do not play a relevant role in the direct transmission; however, they may be important in the indirect transmission of brucellosis. For instance, wild scavengers and dogs may spread contamination by carrying aborted fetuses from infected animals to various sites (Davis, 1990). Accidental hosts, such as humans and horses are frequently infected through the ingestion or inopportune contacts with contaminated raw tissues or milk. From the epidemiological point of view, these accidental hosts are irrelevant for the transmission of Brucella organisms. Very little is known regarding the transmission of Brucella among marine mammals. It is thought that newborns may acquire the bacteria congenitally or through milk from infected mothers. Nematodes of seals have been proposed to play a role in the transmission of Brucella in these animals (Garner et al., 1997).

Vaccines and Vaccination Long experience has shown that vaccination is an essential tool in the eradication of animal brucellosis (Alton, 1977). Since humans contract the disease from animals and are themselves not a source of contagion, animal vaccination is also decisive in eradicating human brucellosis. Although attempts to vaccinate human beings with some of the living animal vaccines or with subcellular vaccines have been made, these practices have been abandoned. Access to the information on human vaccination with live vaccines is difficult to obtain, but it seems that where implemented, vaccination was discontinued because of the lack of attenuation of the vaccine and/or its noxious side effects. There is information on a subcellular vaccine based on the phenol-insoluble fraction of smooth Brucella (Bentejac et al., 1984), but its actual value in humans has never been rigorously tested, and it does not induce protection in suitable animal models (De Bolpe and Garcia-Carrillo, 1991). Although there is a growing interest in developing attenuated strains suitable for human vaccination, none has been tested except in laboratory animals (Crawford et al., 1996; Drazek et al., 1995; Hoover et al., 1999). Protective immunity achieved by Brucella vaccines depends upon several factors, such as animal species, age, gender, administration route, and vaccine dose, type and challenge. Vaccination has been extensively used in cattle, sheep, and goats but not in other domestic

376

E. Moreno and I. Moriyó n

animals, either because the breeding conditions do not make it necessary, or because attempts have been discouraging. None of the existing vaccines against animal brucellosis induce lifelong 100% protection against the challenge encountered in heavily infected flocks, and the most effective vaccines interfere in the serological diagnosis of brucellosis. To solve these problems, subcellular vaccines, adjuvant formulations, and different vaccine strains have been investigated. Moreover, for the classical live vaccines (i.e., B. abortus strain 19 and B. melitensis strain Rev. 1), variations in the dose, route, age of the animals at vaccination and tests used in the serological follow-up have been studied. So far, only these live vaccines have clearly proven prophylactic efficacy in the natural hosts. The use of dead vaccines in adjuvant is generally considered much less effective, and little research on subcellular vaccines has progressed beyond the laboratory animal models (Blasco, 1990; Nicoletti, 1990a). Attenuated mutants derived from the identification of genes related to virulence (Vaccines and Vaccination; Brucella Virulence Mechanisms) have been rarely tested in the natural hosts and either the studies have not been continued (Cheville et al., 1993) or the results are inferior to those obtained with the classical vaccines (Elzer et al., 1998). Animals are generally immunized with the vaccine strain homologous to the Brucella spp. for the corresponding host range (i.e., B. melitensis vaccines are used in sheep and goats and B. abortus in cattle). There have been attempts to use B. abortus strain 19 in small ruminants but the results have been less satisfactory than those obtained with the homologous vaccine. Although preliminary reports are encouraging (GarciaCarrillo, 1980; Horwell and Van Drimmelen, 1971), B. melitensis Rev. 1 has not been used in cattle and its safety in this host is unknown. B. MELITENSIS Rev 1 Attenuated B. melitensis biotype 1 Rev. 1 (revision one) vaccine has been efficiently used for more that 40 years in the prevention of caprine and ovine brucellosis, including B. ovis infections (Blasco et al., 1984b; Blasco et al., 1987; Marín et al., 1999). Although it is not exempt of problems (Origin, Characteristics and Attenuation of Rev 1), vaccination of sheep with the standard dose by the conjunctival route is the most practical means to rapidly reduce the rate of B. melitensis infection in both sheep and humans (Garin-Bastuji et al., 1998). This smooth-strain vaccine displays some residual virulence inducing abortions and testicular problems when administrated to sexually mature caprines and ovines, and it is also capable of infecting humans (Crespo-Leó n, 1994). Some female animals vaccinated with Rev. 1 may

CHAPTER 3.1.16

release the bacteria in milk for several weeks or months. Origin, Characteristics and Attenuation of Rev 1 B. melitensis Rev. 1 is strain number 1 of a set of revertants obtained from a streptomycindependent strain by Elberg and his colleagues at the University of California in the mid-1950s (Alton and Elberg, 1967, 1981). This strain has a smooth phenotype (Response to Environmental Stress), but also has some traits that allow its differentiation from field strains (Identification of Vaccine Strains), an essential property in a live vaccine against brucellosis. Its rate of smoothto-rough dissociation (with the ensuing overattenuation) is relatively high and calls for strict vaccine quality control. Moreover, the biological properties of Rev. 1 seeds conserved in different laboratories seem to have shifted through the years, so that the bacteria in different commercial vaccines may vary in critical properties. There are demonstrated cases in which the particular vaccine used clearly lacked attenuation (Blasco, 1997), whereas in other cases the vaccine varied to the rough phenotype (E. DíazAparicio, personal communication). Biological quality control procedures, which should be implemented at least for periodic seed stock control, have been developed to measure Rev. 1 residual virulence and immunogenicity (Bosseray, 1991; Grillóet al., 2000). The original Rev. 1 strain is attenuated in mice and guinea pigs, where it develops an infection milder than that caused by virulent strains and elicits protective immunity. It is also attenuated in goats and sheep where it induces a generalized infection, which is practically eliminated from most animals by the fourth month post vaccination (Alton and Elberg, 1967, 1981). The cause(s) of the attenuation is not known but the dye sensitivity pattern suggests that at least some properties of the outer-membrane are altered. It is important to stress that, despite its attenuation in animals, Rev. 1 is clearly infectious for humans (Alton and Elberg, 1967, 1981) and resistant to streptomycin (Human Brucellosis). Although it is rarely excreted in sheep milk, goats are reported to excrete Rev. 1 more often and for longer times, thus posing a hazard for farmers (Alton, 1990a). Moreover, veterinarians applying the vaccine and workers restraining the animals during this process are also at risk. Protection of Rev 1 The vaccine can be administered either subcutaneously or conjunctivally with no differences in the degree of protection obtained. In controlled experiments, it has been demonstrated that at the optimal (or standard) dose (about 109 colony forming units [CFUs]) Rev. 1 protects 70–85% of the sheep against a

CHAPTER 3.1.16

challenge (about 108 CFU of a virulent B. melitensis strain) infecting 75–100% of unvaccinated controls (Blasco, 1997; Garin-Bastuji et al., 1998). Under field conditions, the immunity obtained in sheep and goats is satisfactory for eradication purposes when vaccination is combined with the removal of infected animals detected in the most specific serological tests (Alton, 1990a; Garin-Bastuji et al., 1998). Controlled studies in goats show that immunity induced by Rev. 1 vaccination may last longer than four years. In sheep, the experimental evidence shows a high level of immunity for at least two and a half years (Alton, 1990a). However, it is not known whether vaccination of young animals with the standard dose induces lifelong immunity. Obviously, adult vaccination would be desirable for eradication and in fact must be applied when programs start and prevalence is high (Garin-Bastuji et al., 1998), but this increases the side effects of vaccination. Also, Rev 1 is the best vaccine available against B. ovis ram epididymitis, and the conditions of use are the same as in ewes (Blasco, 1990; Marín et al., 1990). Interference in the Serological Diagnosis and Other Side Effects of Rev 1 Because it is a smooth strain, Rev. 1 induces antibodies interfering in the smooth-LPS serological tests and this complicates the diagnosis (Immunological Diagnosis of Animal Brucellosis Caused by Smooth Brucella). The duration of this response depends on the age of the animals at vaccination. The duration is longer as they approach or reach sexual maturity. It is significantly more protracted when the subcutaneous route is used and, in this case, many animals show positive serological results for years. When using the best strategy (i.e., the conjunctival route and less than 6month-old animals), the great majority of the animals become serologically negative in the most specific serological test in less than a year (Alton, 1990a; Fensterbank, 1987; Garin-Bastuji et al., 1998; Jiménez de Bagü és et al., 1989; Plommet and Fensterbank, 1984; Zundel et al., 1992). Rev. 1 is very abortifacient and rates of vaccine-induced abortions can be as high as 80% if animals are vaccinated in the second to third month of pregnancy (Alton, 1990a; Alton and Elberg, 1967; Blasco, 1997; Garin-Bastuji et al., 1998). The risk of inducing abortions is substantially reduced but not eliminated when the conjunctival route of vaccination is used (Blasco, 1997; Jiménez de Bagü és et al., 1989; Zundel et al., 1992). Alternatively, the use of reduced doses (from 105 to 107 CFU) has been proposed but abortions are not totally avoided, and there is clear evidence that the immunity achieved is not adequate (Blasco, 1997).

The Genus Brucella

377

B. ABORTUS Strain 19 B. abortus strain 19 vaccine has been extensively used in cattle. With a few exceptions, most of the successful control and eradication programs have been carried out with this strain. It is noteworthy that this vaccine was developed more than 70 years ago and that, although several alternative attenuated strains have been developed subsequently and studied, none has such a wide application as strain 19. Origin, Characteristics and Attenuation of S19 Strain 19 is derived from the nineteenth stock culture made by Buck from a milk isolate in the early twenties. The culture was maintained at room temperature for over a year and this resulted in attenuation in the guinea pig. Strain 19 has a smooth phenotype (Response to Environmental Stress), and since occasionally it dissociates to yield rough mutants unsuitable for vaccination (Colonies and Dissociation), quality control of at least this property is necessary. It carries some traits that allow its typing and differentiation from field strains (Identification of Vaccine Strains), including inhibition by erythritol (Erythritol). As compared to field isolates, strain 19 is also markedly attenuated in cattle and only induces a transient infection in heifers that is usually cleared in a few months and seldom lasts for more than a year (Nicoletti, 1990a). When used in adults, however, about 2% of the animals develop udder infections and shed the vaccine in the milk for several years. Although the precise reasons for the attenuation are not known, strain 19 clearly shows altered intracellular trafficking (Association with the Autophagic Machinery; Interleukins in Brucellosis). The deletion in the eri region carried by this vaccine and the alteration of outer-membrane properties suggested by its antibiotic and dye sensitivity pattern may be related to the loss of virulence. Although not as dangerous as B. melitensis Rev. 1, B. abortus strain 19 also has been a demonstrated cause of human infections (Meyer, 1985). Protection of S19 In controlled experiments, 40–75% of heifers vaccinated subcutaneously with the standard dose (1011 CFU) are protected against conjunctival challenges infecting 100% (about 107 CFU) to 87% (about 106 CFU) of the unvaccinated controls (Nicoletti, 1990a), and the level of protection is no different when the conjunctival route is used (Nicoletti et al., 1978; Plommet and Fensterbank, 1984). Although these experiments also show that the immunity obtained can be overcome by a heavy challenge, it has been repeatedly shown under field conditions that strain 19 vaccination reduces the prevalence to very low levels and that eradication is possible when the vaccine is combined with the

378

E. Moreno and I. Moriyó n

removal of the reactors in serological tests of the adequate specificity (Immunological Diagnosis of Animal Brucellosis Caused by Smooth Brucella). Protection is reported to last for more than five pregnancies, and it is not clear whether revaccination increases immunity (Nicoletti, 1990a). A single dose has proved to be enough to achieve eradication in test and slaughter programs applied to herds varying from medium to small size. In these cases, the problems created by revaccination in the interpretation of the serological tests (Interference in the Serological Diagnosis and Other Side Effects of S19) would outweigh the potential benefits. On the other hand, revaccination may be necessary to control the disease in large herds, or under conditions in which obligatory slaughtering is not economically viable, as is the case in low income countries (Nicoletti, 1990a). This strategy is the most practical one in those cases where diagnosis is not routinely carried out as part of a control program and where abortion is causing the major losses. Revaccination in these cases slows the abortion rate and decreases the level of infection by lowering the bacterial inocula. From this perspective, revaccination must be considered an emergency strategy. Some experiments have shown that the best protection is obtained by subcutaneous vaccination of calves followed by a conjunctivally administered booster dose (Plommet and Fensterbank, 1984). Interference in the Serological Diagnosis and Other Side Effects of S19 Although the level of protection induced by strain 19 is good, the presence of residual antibodies against the Opolysaccharide complicates the differentiation between vaccinated and infected individuals. To solve this problem, four major strategies have been proposed (Hidalgo et al., 1977): (1) to develop sensitive and specific serological tests capable of differentiating infected from vaccinated animals, (2) to limit vaccination to calves, (3) to reduce the vaccine dose, and (4) to administer the vaccine by the conjunctival route. Each strategy has its benefits and drawbacks. Although the combination of serological tests under defined circumstances may achieve sensitivity and specificity close to 100% (Immunological Diagnosis of Human Brucellosis Caused by Smooth Brucella), the specificity and sensitivity of these assays considerably decrease under conditions in which infection of some vaccinated individuals occurs or when revaccination is used. Calf vaccination shortens the time lapse during which there are significant levels of antibodies against the O-polysaccharide, but the level of protection is lower than that obtained in adult animals (Nicoletti, 1977). The optimal age of calf vaccination is between 3 and 6 months, and

CHAPTER 3.1.16

under these conditions, most animals are negative in the most specific tests a year later. On the other hand, if vaccinated when they are 6 months to one year old, up to 20% of the animals may be serologically positive 18 months later (Nicoletti, 1990a; J. M. Blasco, personal communication). As discussed above, there are a number of situations that make vaccination of adults necessary but the vaccine persists in adults for a longer time than it does in calves and induces a more protracted serological response. Conjunctival vaccination (two doses of 5 × 109 CFU at a 4–6 month interval) dramatically reduces the serological response without compromising the level of immunity obtained (Nicoletti, 1977; Nicoletti et al., 1978; Plommet and Fensterbank, 1984), and reducing the subcutaneous dose (from 5 × 109 to 5 × 108 CFU) also reduces the serological response. In a large field study, no differences in prevalence rates were found between groups of animals that had received the standard or the reduced dose, and the usefulness of the latter has been confirmed in other studies (Nicoletti, 1990a). However, reduced doses have to be used with care and always considering the age of the animals and particular conditions of the herds as other studies have shown that both younger and old heifers are not adequately protected by strain 19 at a dose of 1 × 09 to 9 × 106 CFU (Huber et al., 1990). B. abortus strain 19 is abortifacient when applied to pregnant cattle but the rate of abortions is relatively low (about 5%). As mentioned (Origin, Characteristics and Attenuation of S19), adults may develop udder infections and shed the vaccine in their milk. B. SUIS Strain 2 An attenuated strain (strain 2 or S2) was obtained in China after serial transfer of a virulent B. suis biotype 1 strain on culture media for years. Brucella suis strain 2 is of smooth type, shows the same level of attenuation as strain 19 shows in guinea pigs, and does not revert to the virulent state after repeated passaging through guinea pigs, sheep, goats or boars. It is reported to protect cattle, goats, sheep and pigs when administered orally. Implementation of vaccination programs in semiarid grasslands where sheep and goat breeding is extensive is necessarily difficult. Since in these areas the only source of drinking water is from wells, B. suis strain 2 has been administered orally in the drinking water. By this route, the S2 vaccine has been widely used for prevention of animal brucellosis in China (Xin, 1986). Good results were also obtained with sheep in a trial in Libya (Mustafa and Abusowa, 1993). However, in controlled experiments performed in France (Verger et al., 1995) and Spain (Blasco et al., 1993b), B. suis strain 2 administered subcutaneously did not

CHAPTER 3.1.16

protect sheep against a B. melitensis or a B. ovis challenge under conditions in which the standard B. melitensis Rev. 1 was effective. The different breeds employed, differences in the challenging strains used, the climatic and nutritional conditions and the difficulties in obtaining reliable data under field conditions may account for the discrepancies. Rough Vaccines The use of rough vaccines has been proposed on the premise that protective immunity in bovine brucellosis is carried exclusively by cells and that the generation of antibodies against the O-polysaccharide is unnecessary and undesirable, since it complicates the differential diagnosis (Cunningham, 1977a). B. abortus 45/20 A killed vaccine prepared with the rough strain 45/20 suspended in a water-inmineral oil emulsion has been used in the past (Cunningham, 1977a). This vaccine was developed to overcome the disadvantages of the interference in the serological tests caused by strain 19, which limited its use to calves before the development of more specific serological tests and the demonstration of the usefulness of reduced doses and the conjunctival route in adult cattle. However, strain 45/20 was shown to revert to smooth pathogenic forms when injected into cattle, and this limited its use to a killed in adjuvant vaccine (Adams, 1990; Chukwu, 1985). Although vaccine 45/20 induces some protection in adults, calves of less than six months old may not be well protected (Nicoletti, 1990a). Also, some commercial 45/20 vaccines undoubtedly induced antibodies to the smooth-LPS (Díaz and Jones, 1973b), thus reducing the only advantage of this vaccine. In countries where the conditions were favorable (low prevalence and intensive breeding with good animal management practices), vaccine 45/20 was found to be useful in providing herd resistance (Roerink, 1969) but other experiences have been less favorable (Plenderleith, 1970; Ray, 1976). B. abortus RB51 In recent years, a great deal of attention has been placed on the possibilities that B. abortus strain RB51 offers as a live vaccine (Schurig et al., 1991). Strain RB51 is a spontaneous rough mutant selected after repeated in vitro passage of B. abortus 2308 (United States Department of Agriculture challenge strain) in the presence of rifampin and penicillin (Schurig et al., 1991). Selection was performed using the crystal violet and acriflavine tests (Colonies and Dissociation) and a considerable effort has been dedicated to the genetic characterization of RB51. The RB51 strain carries an IS711 spontaneously inserted in the wboA gene (Table 5), which is undoubtedly related to its phenotype

The Genus Brucella

379

and attenuation (Vemulapalli et al., 1999). However, B. abortus 2308 wboA mutants obtained by transposon mutagenesis are not equivalent to RB51, which shows that the latter strain carries additional and unknown defects (McQuiston et al., 1999). The strain is stable and, although it should show very little or no virulence in humans, there is little experience on this point. RB51 infections cannot be diagnosed in the standard serological test, as this vaccine lacks the O-polysaccharide (see Immunological Tests for Brucellosis), and it has to be stressed that it is resistant to rifampin (Treatment). Being a rough strain, RB51, although it produces small quantities of N-formylperosamine polysaccharides (G. G. Schurig, personal communication), does not induce antibodies interfering in the smooth-LPS serological tests (Stevens et al., 1994b) even when used as a booster in adult animals vaccinated with strain 19 during calfhood (Uza et al., 2000). This vaccine has been reported not to be abortifacient in cattle (Edmonds et al., 1999; Palmer et al., 1997; Uza et al., 2000). However, animals vaccinated at pregnancy excrete it in the milk (Uza et al., 2000) and full doses of RB51 (1 × 1011 CFU) administered intravenously induce severe placentitis and placental infection (Palmer et al., 1996). Moreover, recent evaluations of some field experiences have demonstrated high numbers of RB51 isolates in aborted cattle (Lopetegui, 1999). When using the reduced dose of this vaccine (1 × 109 CFU), no abortions or placentitis lesions are produced in subcutaneously vaccinated cattle (Palmer et al., 1997). However, this reduced dose does not protect against B. abortus (Olsen, 2000). The data on the level of protection afforded by RB51 in cattle are contradictory. In field studies carried out in Venezuela, it was concluded that RB51 was superior to strain 19 (both used at a dose of 5 × 109 CFU) in protecting against the natural challenge existing in infected herds (Lord et al., 1998b). On the other hand, in a controlled experiment, RB51 protected 26 of 29 vaccinated animals and strain 19 protected 20 of 22 (both used at a dose of 1010 CFU), but only a moderate challenge was used in the experiment (12 of 20 nonvaccinated controls were infected) (Cheville et al., 1996b). More recently, the results of a study in which 3-month-old heifers were given RB51 at doses ranging from 109 to 1.4 × 1011 have been presented (Olsen, 2000). Four of eight nonvaccinates, two of four 109 CFU vaccinates, and two of fourteen 1.4 × 1011 CFU vaccinates aborted. In a separate study conducted with 6-month-old heifers, three of seven nonvaccinates and one of eighteen 1010 CFU vaccinates aborted. The author concluded that RB51 was efficacious in preventing abortion and fetal infec-

380

E. Moreno and I. Moriyó n

tions, but the data also show that the incidence of maternal infection with Brucella at necropsy did not differ among RB51 vaccinates and nonvaccinated controls (Olsen, 2000). Additional comparative experiments with strain 19 under controlled conditions are necessary to define the level of protection achieved with this vaccine in cattle, and it is still premature to draw conclusions on its ability to protect under field conditions. Up to now, RB51 vaccine seems to prevent brucellosis under conditions of low prevalence; however, under conditions of high prevalence, this vaccine has not been satisfactorily tested. Our own experience is that under field conditions, strain 19 performs better than RB51, at least in circumstances where high inocula of virulent Brucella organisms (high incidence in the herd) are likely to occur (E. Moreno et al., unpublished observations). B. abortus RB51 does not seem useful in sheep. In a stringent experiment (100% of infection in unvaccinated controls), RB51 afforded no protection against B. ovis under conditions in which Rev. 1 fully protected 40% of the animals (Jiménez de Bagü és et al., 1995). Moreover, controlled evaluations against B. melitensis challenges are also unsatisfactory (el-Idrissi et al., 2001). There is also one study in pigs, although conducted under field conditions. In these animals, the vaccine was reported to induce 100% protection regardless of the dose (106 or 109), the number of injections (1 or 3) and the administration route (oral and intramuscular) (Lord et al., 1998a). Controlled experiments would be necessary for a further evaluation of the vaccine in pigs.

Control, Eradication and Prevention Control and subsequent eradication of brucellosis can be consummated by the simple, costly and sometimes impractical procedure of totally eliminating the primary hosts from the designated area, with or without replacement of the population with clean animals. Although this method has been attempted by some, most countries that have achieved eradication of brucellosis from bovine, ovine and caprine herds have followed a stepwise strategy (Boyd, 1977; Flowers, 1977). The first initiative towards the control of brucellosis in a specific population usually consists of the serological examination of the animals (Immunological Tests for Brucellosis). Under circumstances where the animal population has not been vaccinated, a simple sensitive test (i.e., card or rose bengal) is enough for identification of the infected animals. However, when strain 19 vaccination is carried out, a combination of sensitive and highly specific assays (Immunological Diagnosis of Animal Brucellosis Caused by

CHAPTER 3.1.16

Smooth Brucella) for the differentiation of infected and vaccinated animals (i.e., complement fixation, radial immunodiffusion [RID], enzyme-linked immunoabsorbent assay [ELISA] or competitive ELISA) is required. The second initiative is the vaccination of the population at risk followed by removal and slaughtering of the serologically positive animals. During this phase, brucellosis-free farms and regions with or without vaccination could be declared. The final step is to stop vaccination. Constant serological surveys are necessary, mainly when potential foci of contamination are colliding with brucellosis-free areas. Only after several years of continuous surveillance, brucellosis may be considered eradicated from these nonvaccinated serologically negative herds. Up to now, only strain 19 and Rev. 1 vaccines have demonstrated to control and eradicate brucellosis under conditions where control programs have been implemented (Blasco, 1997; Nicoletti, 1990a). One very important aspect in the prevention of brucellosis outbreaks is to limit and strictly control the import of animals from regions where brucellosis has not been eradicated. Similar to this is the introduction of susceptible hosts into areas where brucellosis may remain latent in the population. This is especially important in cases involving wildlife hosts, such as the introduction of bovines, goats or ovines in regions where bisons, reindeer and perhaps marine mammals remain as potential reservoirs of Brucella organisms.

Zoonosis Human brucellosis is a highly contagious bacterial zoonosis, which follows the distribution of the natural animal hosts (Fig. 29). Removal of the infected hosts, therefore, eliminates the contagion risk. Human brucellosis is most frequently found in third world countries, mainly due to the high prevalence of Brucella infections in domestic herds and to the ingestion of unpasteurized dairy products. The disease in humans is also known as “undulant fever” or “Malta fever,” since the first isolation of the causative bacterium was made in the Mediterranean island of Malta from spleens of British soldiers who became infected as a consequence of drinking contaminated caprine milk. The most common Brucella species infecting humans are B. melitensis, B. abortus and B. suis. Among these, B. melitensis and B. suis are the most aggressive species for humans. These bacteria cause a severe syndrome, which if not treated may lead to death. B. canis has been reported sporadically to infect humans. B. ovis, B. neotomae and the marine Brucella strains have not been detected in humans. The marine

CHAPTER 3.1.16

strains may represent, however, an authentic hazard for human communities that hunt and gain subsistence from whales and seals, as well as for workers that are in close contact with marine mammals (Garner et al., 1997; Higgins, 2000). Contamination of humans depends upon interaction with infected animals, their products, or their secretions. Because of the wide acceptance of pasteurization, brucellosis is regarded in many countries as an occupational disease of sporadic occurrence among livestock handlers, veterinarians, vaccinators, laboratory workers, and meat-processing and slaughter plant personnel. In many third world countries the consumption of raw milk and unpasteurized dairy products, such as butter, fresh cheese, cream or curd, as well as fresh raw viscera and blood is still an important source of infection. The infection of domestic animals, frequently accompanied by slaughtering, and the possibility of contamination of food products with Brucella add important restrictions to the marketing of foodstuffs. In consequence, incomes for many families are reduced. The diagnosis of human brucellosis is well established (Antibody Detecting Tests in Infected Patients). It depends mainly on the detection of antibodies in the serum of patients against Brucella organisms by immunological techniques. The confirmation of infection is achieved by the isolation of the etiological agent in the blood of infected patients. Although pasteurization is the mandatory public health procedure, examination of foodstuffs, presumed to be the source of contagion, may occasionally be necessary. The demonstration of antibodies in milk is suggestive of herd infection and of the possible presence of Brucella spp. in dairy products, but it is not a specific method of diagnosis because revaccination or adult vaccination can cause secretion of antibodies in milk. Shedding of the vaccine strains by the immunized animals may also occur. Since vaccine strains are virulent for humans, their presence in dairy products constitutes a zoonotic risk. Isolation of Brucella organisms can be achieved from milk, but it is more difficult when processed dairy products are concerned, complicating matters even more. Culture on selective media follows the same procedures described for animal diagnosis by bacteriological culture (Molecular Tests for Humans; Molecular Tests for Animals). Molecular methods, such as PCR, may be useful and could be used to detect some of the vaccine strains (Molecular Tests for Humans; Molecular Tests for Animals), but there are few reported studies on their application to processed foodstuffs.

The Genus Brucella

381

Disease Brucellosis is a disease of the reticuloendothelial system, the reproductive organs and of the fetus, and then the pathological signs are perceptible accordingly (Pathology). In the primary host the main manifestation of the disease is abortion or epididymitis. In humans the syndrome is in extreme complex and could be evidenced in many forms, recurrent fever being one of the main manifestations. From a gross perspective, the pathobiology of brucellosis is in several respects similar to that of other intracellular bacteria; however, it possesses many idiosyncratic manifestations and a unique intracellular life cycle (Pathobiology). The infection is controlled by an efficient immune cellular response (Control of the Infection), although antibiotic therapy is mandatory in humans (Treatment). Diagnosis is mainly based in determining the immune response of the infected host and in the isolation of the bacteria from infected tissues and blood (Diagnosis).

Pathology The pathology of brucellosis is complex and its study requires biosafety conditions (Biosafety) due to the fact that the bacteria are primary pathogens of animals (Brucellosis in the Host Animal) and humans (Brucellosis in Humans) and then are a risk for contamination. For comprehensive reviews on the pathology of brucellosis, the publications of Blasco (1990), Enright (1990a) and Spink (1956) are recommended. The Disease Named Brucellosis Considering the close phylogenetic relationship among the different members of the genus, it is not surprising that the pathologies induced by the various Brucella species in their host animals are very similar. Likewise, the gross physiopathological events that occur in accidental or experimental infections in secondary hosts, such as humans, mice and horses, have many things in common. Although some strains are more virulent than others, the different pathologies observed seem to be related more to idiosyncratic differences of the hosts than to individual virulence properties of the Brucella strains. This means that once Brucella organisms have gained access into the tissues, they proceed in a very similar manner. Control of the infection depends mostly on the confrontation between the host defenses, on one side, and the ability of Brucella to reproduce and gain access to various tissues without being destroyed, on the other (Control of the Infection). Most of the pathobiological studies have been performed in experimental animals, such as mice

382

E. Moreno and I. Moriyó n

and guinea pigs, as well as with a variety of cell lines. In the natural hosts, studies have been conducted with bovines, caprines and ovines and with strains of B. abortus, B. melitensis and B. suis. Although the data are more limited, similar pathologies have been observed with the marine strains B. “maris,” B. canis and B. ovis in their natural hosts. This last species, however, demonstrates preference for male ovine reproductive organs, with low pathogenicity for ewes (Blasco, 1990). Similar to B. ovis, the wood rat parasite B. neotomae seems to display low virulence, even though production of highly aggressive strains generating similar events to other Brucella species has been reported (Gibby and Gibby, 1965). Finally, the study of human brucellosis has contributed to the understanding of the pathology and pathobiology from various perspectives. Brucellosis in the Host Animal In general terms, brucellosis can be viewed as a chronic infection, which in nontreated cases may cure or persist for life, mainly within cells of the reticuloendothelial system (Blasco, 1990; Enright, 1990b). The incubation period in the preferred animal host under natural conditions depends upon the Brucella strain, inoculum size, as well as upon host factors. In the natural host, incubation times range from two to five weeks, although longer periods are not exceptional. In the accidental host, the incubation period is more variable and generally longer than in the natural host animal, often lasting for several months. The onset of symptoms in the natural hosts normally courses with recurrent fever and enlarged lymph nodes, spleen and liver. Arthritis with gastrointestinal and nervous symptoms may be observed. Bacteremia normally coincides with the fever events. In the natural pregnant host, the infection frequently results in abortion in the last trimester of gestation and in contagion to other members of the herd. In males, invasion of the reproductive organs is frequently observed, facilitating venereal transmission and inducing partial or total sterility. Orchitis and epididymitis are two prevailing signs. In the immunocompromised host, invasion of many organs, including the central nervous system, may take place followed by death. Reproductive impairment in females may occur upon placental retention or for other causes; however, this event is exceptional rather than a common outcome in brucellosis. The most frequent ports of Brucella entrance are the mucous membranes of the oral, respiratory and gastrointestinal tracts, as well as those membranes covering the conjunctiva, vagina and the prepuces (Enright, 1990b). Very likely bacterial penetration takes place through the epithelial cells lining the mucous membranes (Brucella

CHAPTER 3.1.16

Invades Healthy Hosts). At the site of lodgment, the organisms are ingested by resident phagocytic cells and then carried via the lymphatics to the regional lymph nodes. Once ingested, the bacteria travel to a specialized intracellular compartment where they multiply. Some of the bacteria and cells die during the confrontation, and the released Brucella debris stimulate a local immune response with the concomitant activation and proliferation of inflammatory and mononuclear cells (Anderson et al., 1986b; Anderson et al., 1986c; Cheville et al., 1992; Tobias, 1993). The result of this confrontation determines whether the infection is contained or transgresses to other organs. In the immunized host, the memory and activated cells of the immune system normally are capable of dominating the infection (Control of the Infection). If not, polymorphonuclear leukocytes and macrophages carrying the bacteria reach the blood, transporting the microorganisms to the reticuloendothelial system, mammary glands and reproductive organs (Enright, 1990b). The histopathological signs of the colonized lymph nodes are typical of infections by intracellular bacteria, demonstrating hyperplasia with mononuclear infiltration, local hemorrhage, presence of granulomas with histiocytes, epithelioid cells and gigantic Langerhans cells, edema and extramedullary hematopoiesis. Severe paracortical lymphocyte depletion, destruction of germinal centers and serofibrous lymphadenitis with effacement of lymph node architecture are observed during the first weeks of infection. In most cases, dead and alive intracellular bacteria can be detected in large numbers within phagocytic cells and in lesser proportion within lymphocytes (Fig. 30). The Fetus, the Most Susceptible Host In pregnant animals, such as bovines, caprines, ovines, swine and very probably marine mammals, the bacteria reach the gravid uterus, infecting the cotyledons (Anderson et al., 1986b; Anderson et al., 1986c). Once installed in this site, the bacteria replicate within the endoplasmic reticulum of erythrophagocytic trophoblasts followed by extensive replication within chorioallantoic trophoblasts (Fig. 30). This action generates necrosis and subsequent ulceration of the chorioallantoic membrane with production of anti-inflammatory exudates composed of phagocytic cells and cellular debris (Fig. 31). At this point, Brucella can be observed within the uterine lumen and in the placental capillaries disseminating through the chorionic villi. As a consequence, placentomal and fetal infection with concomitant inflammation proceeds. Interestingly, Brucella organisms are not found within placentomal trophoblasts, cells of the

CHAPTER 3.1.16

The Genus Brucella

A

B

C

D

E

F

383

Lym

Fig. 30. Intracellular parasitism of Brucella in different host cells. Transit of B. abortus in caprine M (lymphoepithelial) cells (A); dividing HeLa cell with replicating B. abortus (in red) within the endoplasmic reticulum (ER) (immunofluorescence) (B); caprine trophoblast with replicating Brucella within the ER (C); human polymorphonuclear leukocyte with intracellular B. abortus in phagosomes and phagolysosomes (D); murine macrophage with replicating B. abortus within cisterna resembling endoplasmic reticulum (E); and caprine lymphocyte with intravacuolar B. melitensis (F). In (A), extracellular material was co-ingested with Brucella organisms. Notice in (B, C and E) that the cell nucleus is constrained but not invaded by the replicating brucellae. Notice in (B) that karyokinesis is not inhibited by the replicating brucellae. Notice in (D) that several azurophil granules fuse with phagosomes containing Brucella. In spite of the large number of intracellular bacteria the cells do not present signs of apoptosis or necrosis. The arrows indicate a single bacterium within a vacuolar compartment. Panel (A) is from Ackermann et al. (1988), panel (C) from Anderson et al. (1986b), panel (D) from Riley and Robertson (1984a), panel (E) from Jiang and Baldwin (1993b), and panel (F) from Cheville at al. (1996a), all with permission.

endometrium or syncytial epithelium (Fig. 32). Abortion without significant compromise and bacterial invasion of the endometrium or lamina propia are characteristic of this phase. Extensive necrosis promotes diffusion of Brucella into different adjacent tissues. Vasculitis with separation of the maternal epithelium promotes fetal death and abortion. Ulcerative endometritis and metritis may occur as a consequence of postparturient infections with opportunistic bacteria. From the clinical point of view, abortion due to brucellosis resembles other infectious diseases, although

retention of placental tissues or endometritis seldom occurs in the natural hosts. Following Brucella invasion and depending on the age of the fetus, leukocytosis with significant numbers of neutrophils develops. In ruminants, the infected fetuses demonstrate lymphoreticular hyperplasia of secondary lymphatic organs and extensive granulomatosus centers composed of macrophages, histiocytes, epithelioid cells, gigantic Langerhans cells, extramedullary hematopoiesis and a variable number of polymorphonuclear granulocytes. Liquefaction and

384

E. Moreno and I. Moriyó n

CHAPTER 3.1.16 Br

Tr

C

B

Br A

Tr

CA

S

P U

C

D

Fig. 31. Pathological examination of placentoma from a pregnant goat infected with B. abortus. A) Diffuse chorioallantoic edema (arrows) obscuring view of the placentoma in a goat given 109 B. abortus cells intravenously and necropsied on postinoculation day 19. White periplacentomal exudate is predominant (arrows). Bar = 1 cm. B) Multifocal abscesses and bacterial colonies of B. abortus in the placentoma are observed in the connective tissue of chorionic villi and rimmed by intact trophoblastic epithelium (tr). Exudate (arrows) separates maternal septa (s) and capillaries (c) from the bacteria-filled chorionic villus. Bar = 30 µm. C) Placentoma from a goat that was inoculated in a middle uterine artery with 109 B. abortus cells and euthanized while aborting on postinoculation day 14. Multiple abscesses and hemorrhages are present in placentomal cross-sections. Uterine wall (u), placentoma (p), chorionic membrane (ca). Bar = 1 cm. Multifocal abscesses and bacterial colonies of B. abortus are observed in the placentoma. D) Bacteria-filled phagocytes (arrow) and necrotic cell debris separating maternal septa (s) from trophoblast (tr). Brown-Hopps Gram stain. Bar = 20 µm. From Anderson et al. (1986a), with permission. ALLANTOIC CAVITY 1 ET COTYLEDON SE

2 PT

CAT CV

3

MS CARUNCLE

4

ALLANTOIC EPITHELIUM CHORIOALLANTOIC MEM. UTERINE LUMEN

TO FELUS UL

ENDOMETRIUM

Fig. 32. Diagram summarizing the pathogenesis of placental and fetal infection in a caprine placentoma, as determined in this study. 1) B. abortus first seen in phagosomes of erythrophagocytic trophoblasts (ET). 2) Subsequently, B. abortus replicates in the rough endoplasmic reticulum of chorioallantoic trophoblasts (CAT). 3) After trophoblast infection, necrosis of chorioallantoic trophoblasts, spewing of B. abortus into the uterine lumen (UL), and ulceration of the chorioallantoic membrane occur. 4) Coincidentally, intravascular B. abortus, present in placental capillaries, spreads to chorionic villi (CV) and fetal viscera. Brucella abortus is not present in placentomal trophoblast (PT), maternal septa (MS), syncytial epithelium (SE), or endometrium. Erythrophagocytic, placentomal, and chorioallantoic trophoblasts form the continuous epithelial sheet that covers chorionic villi of the cotyledon and the chorioallantoic membrane. They are differentiated here, owing to differences in anatomic location, function, and role in Brucella placentitis. From Anderson et al. (1986b), with permission.

CHAPTER 3.1.16

mineralization of the necrotic zones may occur. Commonly, these reactions are also observed in a variety of fetal organs, such as the liver, lungs and kidney. Brucella invasion of the central nervous system is a frequent outcome. Male Infections In male ruminants, bilateral acute orchitis with extensive irreversible damage is a frequent event (Blasco, 1990; Enright, 1990b). After invasion and bacterial replication, the tunics become distended, and necrotic foci are evident within the testicular parenchyma, with hemorrhagic foci and purulent exudate. Focal necrotizing epididymitis and spermatic granulomas are frequently observed after the initial inflammatory reactions. Seminal vesiculitis and prostatitis are not uncommon. After these episodes caused by Brucella infection of the male reproductive tract, partial or total sterility remains as a consequence of the chronic infection. Perivascular edema and infiltration of mononuclear cells and polymorphonuclear leukocytes accompany the initial localization into the tubular tissue. Subsequently, the inflamed tubular epithelium develops papillary hyperplasia and local hydropic degeneration and intraepithelial cysts. Eventual destruction of the epithelium, either by bacteria or by inflammatory reaction, leads to extravasation of the spermatozoa. Complete blockage of the epididymis and testicular degeneration with fibrosis represent the outcome of the chronic infection. Damage of epidydimis and prostate epithelial cells acts as a trigger for the production of autoantibodies to spermatozoa, contributing to the general pathology observed in male organs and one of the mechanisms for causing sterility (Serikawa et al., 1984). Brucella ovis is an important cause of epididymitis in rams, and it is less frequently associated with abortions in ewes. Although female ovines are more resistant to the infection than males are, they are important sources of passive venereal transmission to the copulating males. The target organ of B. ovis is the epididymis and accessory sexual glands. The pathological signs caused in ram epididymis are similar to those described above. Brucellosis in Humans Although many of the pathological signs of Brucella infection in the natural hosts are reproduced in humans, idiosyncratic differences are found (Spink, 1956; Young, 1983). The outcome of the disease in humans depends on host factors, inoculum size and the Brucella strain. In general terms, the most aggressive species for humans is B. melitensis, followed by B. suis and B. abortus, although this tendency is not strictly lined up. For instance, B. abortus biotype 5 and B. suis biotype 2 seem to harbor low pathogenicity for man. Limited

The Genus Brucella

385

infections have been described with B. canis. No human infections caused by B. ovis, B. neotomae or B. maris have been reported. However, human brucellosis caused by the marine strains has been barely explored, owing to the fact that these strains were recently described (Higgins, 2000; Jahans et al., 1997). The evolution of brucellosis in humans has the tendency to be more dramatic than in the animal reservoir, with no pathognomonic signs or symptoms. The incubation period is highly variable, with the most common interval between one and three weeks. However, longer periods are not uncommon. After invasion of host tissues, the organism multiplies in regional lymph nodes draining the site of entry and gives rise to a bacteremia, which may vary from short duration to intermittent or prolonged times. Normally this event is accompanied by symptoms that resemble in some aspects a strong cold, with clinical features that include undulant fever, chills, weakness, malaise, sweating, headache, backache, muscle joint pain and anorexia. After this period, and following colonization of the reticuloendothelial system, more severe symptoms may arise which cause weight loss, nausea, cough, vomiting, diarrhea, epistaxis and rash. After this, lymphoadenopathy, hepatomegaly, splenomegaly, osteomyelitis, thrombosis and valvular endocarditis may arise. In many cases compromise of the central and autonomous nervous systems may appear as well as compromise of the urogenital organs. Abortion due to Brucella infections, with invasion of maternal and fetal tissues, has been reported in a limited number of cases. If nontreated, complications due to Brucella occur in about 10% of the cases. These generally involve suppurative abscesses in different organs and tissues. Mortality usually is less than 2%, mainly reduced by antibiotic therapy (Animal Brucellosis). Chronicity with sporadic appearance of symptoms may remain for years. Allergic hypersensitivity is a common consequence of human and animal brucellosis (Díaz and Oyeledum, 1977; Spink, 1956). This is especially important in laboratory workers or medical personnel who may be exposed to Brucella organisms or their products (antigens) on a routine basis. Brucellosis in Experimental Animals In addition to the natural hosts, a number of experimental animal models have been tested including chick embryos, rabbits, rats, hamsters, gerbils, guinea pigs and mice (Garcia-Carrillo, 1990). For many years guinea pigs remained as the preferred model, mainly due to their high susceptibility to Brucella infections and the resemblance of the disease with humans and also because these animals were originally used for

386

E. Moreno and I. Moriyó n

isolating Brucella organisms from clinical samples. In the last 20 years, guinea pigs have been abandoned, and mice have been established as the preferred animal model. The reasons lay mainly in the economical feasibility of the mouse model, the extensive number of available defined strains, the amenability of mice to genetic manipulation, and the extensive knowledge of the immune system of this murine species. However, mice display a relatively strong innate resistance to brucellosis and may not be the more appropriate model for Brucella studies. Brucellosis in Guinea Pigs In guinea pigs, as few as 50 intraperitoneal or intravenously injected Brucella colony forming units (CFU) are required to cause infections, with incubation periods that range from one to three weeks (Ne’eman and Jones, 1963). Although these rodents are susceptible to being infected by oral, conjunctival or respiratory routes, which are estimated to be the most common natural course of infection, safety has prevailed and intraperitoneal, subcutaneous and intramuscular routes are preferred. No major differences have been found between male and female or the various strains of guinea pigs. However, pregnancy seems to be a propensity factor for susceptibility to Brucella infection and is an experimental model to demonstrate vertical transmission and abortion. The initial stages of the infection are similar to those described for other animals, including humans (Moulton and Meyer, 1958; Sterba, 1984). After this initial outcome, splenomegaly, hepatomegaly and compromise of the lymphatic tissues are almost invariably observed. Acute and subacute inflammation occurs with exudation of fluids and accumulation of polymorphonuclear leukocytes. As the infection continues, typical granulomatous lesions with the presence of macrophages, histiocytes, epithelioid cells and giant cells surrounded by proliferating lymphocytes occur. In later stages of the infection, fibrosis is detected (Moulton and Meyer, 1958; Sterba, 1984). In males, compromise of urogenital organs can be observed. In pregnant females, compromise of placental and fetal tissue is evident. Although Brucella organisms have been detected in the semen and fluids of infected guinea pigs, no horizontal transmission has been demonstrated. After several weeks, the infected guinea pigs demonstrate strong delayed-type hypersensitivity reactions when tested with Brucella protein extracts. Brucellosis in Mice It is estimated that, in general, mice are from 103 to 106 times more resistant than guinea pigs to Brucella infections (Taran and Rybasov, 1972). After intravenous inoculation, Brucella organisms are cleared by

CHAPTER 3.1.16

phagocytic cells of the reticuloendothelial system where the bacteria replicate (Cheers, 1984). In the case of intraperitoneal inoculation, Brucella organisms are rapidly cleared by resident phagocytic cells that transport the bacteria to the regional lymph nodes, spleen and liver. Normally during the first stages of the infection, lasting from one to two weeks, there are no manifestations of symptomatic acute brucellosis. However, depending on the size of the inoculum and virulence of the Brucella strain, a variable number of organisms are recovered from the spleen, liver and lymph nodes. Following this, replication of Brucella rapidly increases until a plateau is reached. This period commonly lasts a few days; then, the number of organisms declines until equilibrium has been established, corresponding to the development of the immune response. In spite of the fact that splenomegaly is maintained and the organisms may be isolated up to three months after the experimental infection, no severe pathological symptoms are developed. Histopathological studies during the replicating and steady phases reveal a focus of chronic inflammatory reactions in the spleen, lymph nodes and liver. If large quantities of bacteria are inoculated (>106 per mouse), overloading the innate and immune capabilities, or immunocompromised (hormone- or mucine-treated or genetically immunodeficient) mice are used, granulomas and suppurative infections in different organs are observed, followed by death. In cases where low and moderate numbers of Brucella are administered, limited placental colonization and a few vertical bacterial transmissions to the newborn are observed (Bosseray, 1983; Bosseray, 1984). However, injection of large numbers of Brucella in pregnant mice results in preferential bacterial replication within placental giant trophoblasts and within the visceral yolk sac endoderm. Despite the anatomical differences in placentation between ruminant and rodent (epithelial versus hemochorial) placental damage, fetal infection and fetal death occurs (Tobias et al., 1993), but in contrast to what is observed in the natural hosts, pregnant mice are relatively more resistant to abortion and fetal death. Outcome of the Disease and Self Cure In the natural host, the immune system generally gains control of the infection a few weeks after the appearance of the symptoms (Enright, 1990b; Nicoletti and Winter, 1990b). However, some bacteria may remain in reproductive organs, mammary gland, bones and joins for prolonged periods. Some of the primary infected hosts, therefore, may remain as healthy shedders and of epidemiological importance. In contrast, in the accidental hosts, the bacteria are dispersed

CHAPTER 3.1.16

into different tissues, causing a severe disease with no shedding and in consequence of medical but not epidemiological importance (Alton, 1990a, 1990b; Spink, 1956). In nontreated accidental hosts, such as horses or humans, hepatic involvement in the form of acute diffuse hepatitis with focal necrosis, generation of granulomas, and the compromise of lymphatic organs is a common event of chronicity. Both symptomatic and asymptomatic animals can transmit brucellosis, which means that many of the infected individuals shed the bacteria independently of being ill or healthy. After abortion, infection commonly recedes and self-cure proceeds, which is the most frequent outcome in the natural host (Enright, 1990a). Some bovines are naturally resistant to brucellosis, which is related to the ability of their macrophages to kill Brucella organisms intracellularly (Price et al., 1990; Templeton and Adams, 1990). Moreover, calves, kids and lambs may remain without clinical signs, in spite of drinking milk containing millions of Brucella secreted in the udder by their infected mothers. A low proportion of these infected animals at a young age may abort later in life; however, many remain infected but asymptomatic, demonstrating a good equilibrium between the Brucella invader and the host. Furthermore, several of the infected animals which acquired Brucella by vertical transmission, in addition to being asymptomatic, do not display positive serology (Fig. 9), suggesting anergic immune responses. Other individuals, mainly secondary hosts, may remain allergic for life after cured (Spink, 1956).

Pathobiology After invasion of host tissues, Brucella binds and penetrates cells. Once inside, the bacterium is protected from extracellular microbicidal substances and the new life cycle begins. Intracellularly, Brucella traffics and replicates within vacuolar compartments, avoiding digestive mechanisms of cells by means of virulence factors that allow the bacterium to control its cellular environment for its own benefit. The complexity of these bacterial mechanisms and the cross-talk between the host cell on one side and the Brucella parasite on the other are just being unrevealed. Comprehensive reviews on these topics can be found elsewhere (Baldwin and Winter, 1994; Liautard et al., 1996; PizarroCerdá et al., 1997; Pizarro-Cerdá et al., 1999b; Pizarro-Cerdá et al., 2000). BRUCELLA Invades Healthy Hosts It is known that some bacterial pathogens rapidly gain access to the internal host tissues by penetrating specialized epithelial cells. The penetration of inva-

The Genus Brucella

387

sive bacteria requires the interaction of specific receptors and ligands between the animal and bacterial cells that trigger the internalization and the transference of the pathogen inside the body (Van Nhieu and Sansonetti, 2000). The primary routes of Brucella infection are the cells of mucosal surfaces (Enright, 1990b). In goats, B. abortus laying in the ileum is ingested by M cells using a zipper-like mechanism but not by enterocytes (Ackermann et al., 1988; Fig. 33). In these phagocytic cells, the ingested bacterial organisms are located as single entities or in small clusters, within transient vacuoles that do not show signs of lysosomal fusion. The phagocytic vacuolar membrane does not associate with ribosomes, and some of the Brucella-containing compartments demonstrate coingested opaque material. The engulfed Brucella are eventually translocated by M cells to the gastrointestinalassociated lymphatic space, where they are taken up by macrophages and neutrophils (Ackermann et al., 1988; Fig. 33). The numbers of intracellular Brucella in M cells decrease with time after bacterial inoculation, indicating that microorganisms are steadily translocated. After penetrating mucosal layers, both virulent and vaccine B. abortus strain 19 strains induce an inflammatory response in the submucosa (Cheville et al., 1992). It has been demonstrated that urease-negative B. abortus mutants are virulent when injected intraperitoneally in mice (Grillóet al., 1997). Wild type and urease-deficient mutants also show the same sensitivity to acid. However, when urease-deficient mutants are administered by the oral route, the numbers of bacteria recovered in the intestine or in the spleen are considerably lower than those of the wild-type strain. As in the case of Ureaplasma or Helicobacter pylori, in which urease plays a relevant role in the colonization of the host cells (Cover et al., 1991), it may be that in Brucella this enzyme also participates in modifying the immediate local acid environment (Salyers and Whitt, 1994; Sangari et al., 2000). This is an important aspect that must be taken into consideration because the intestine seems to be the most frequent route for invading the primary and secondary hosts through ingestion of milk and dairy products as well as through licking of the aborted fetus or contaminated secretions. Survival Outside Host Cells Brucella are organisms whose ultimate goal is to propagate inside cells (Habitat). Therefore, when released to the extracellular lumen, these pathogens are deprived of their shelter and natural replicative niche and subjected to the attack of a variety of bactericidal substances present in the plasma and body fluids, against which they must defend. The blood, pulmonary secretion, tears, intestinal flu-

388

E. Moreno and I. Moriyó n

CHAPTER 3.1.16

A

C

M cell

L

Mφ

B

PMN

Fig. 33. Brucella transit in M (lymphoepithelial) cells. A) Dome lymphoepithelial cell from calf ileum, two hr postinoculation. Apical extensions engulf two B. abortus cells labeled with immunogold (arrows). Labeled B. abortus cells in phagolysosome (arrow). Bar = 1 µm. B) Immunogold-labeled B. abortus engulfed by lymphoepithelial cell apical extensions (image is [A] at higher magnification). Bar = 0.2 µm. C) Brucella is found within macrophages (M) and neutrophils (PMN) and also between the base (arrows) of M cells from ileum cells (L) at two hr postinoculation. Abundant apical mitochondria are seen in the M cells (arrow head). From Ackermann et al. (1988), with permission.

ids, milk and colostrum contain substances that participate in protecting the body against bacterial invasion (Opsonization and Complement Susceptibility). In addition, Brucella discharged in the inflammation site is exposed to the local attack of molecules released by leukocytes, such as those present in the azurophilic and secondary granules. In general, it has been demonstrated that Brucella is capable of resisting the microbicidal action of soluble substances released by cells (Modulation of Neutrophil Function and Resistance of Bactericidal Secretions). Despite that, there is not a complete picture of how this is achieved. Various properties, mainly related to the structure and function of the Brucella outer-membrane, have been implicated in the resistance of Brucella to humoral substances (Properties of the Outer Membrane). Opsonization and Complement Susceptibility Work carried out mostly with B. abortus, B. melitensis and B. suis has identified several features of these bacteria which are relevant for parasitism (Brucella Virulence Mechanisms). It is becoming clear that some of these bacterial

properties are modulated by the interaction between antibodies and complement with the Brucella cell surface molecules. Indeed, opsonized bacteria stimulate the oxidative burst and the intracellular killing mechanisms in a higher proportion than that in nonopsonized Brucella (Caron et al., 1994b; Harmon et al., 1987; Harmon et al., 1988; Jiang and Baldwin, 1993a; Jiang et al., 1993c; Kreutzer et al., 1979b; Young et al., 1985). The O-polysaccharide makes B. abortus more resistant to complement-mediated killing in the presence of O-polysaccharidespecific antibody (Corbeil et al., 1988). It is feasible to hypothesize that evasion of opsonization and complement lysis constitute adaptive survival mechanisms employed by Brucella organisms. Animals infected with smooth Brucella characteristically generate a strong antibody response against smooth LPS and NH polysaccharides (Díaz-Aparicio et al., 1993; Marín et al., 1999; Fig. 19). The NHs are LPSindependent molecules intertwined with the Opolysaccharide in the outer-membrane (Aragó n et al., 1996b; Fig. 20). Part of their function is to increase the density of O-type polysaccharides

CHAPTER 3.1.16

Modulation of Neutrophil Function and Resistance of Bactericidal Secretions Although Brucella organisms are moderately resistant to acid (Teixeira-Gomes et al., 2000), they display a strong resistance to bactericidal proteins nor-

389

Infected bovines 24

Number of sera

18

12

6

0 LPS

NH

R-LPS

Lipid A

Vaccinated bovines

24

B. abortus 18 Number of sera

on the cell surface, creating a protective layer that prevents the binding of antibodies and the interaction of lytic complement with the deeper structures of the outer-membrane (Aragó n, 1996a; Fig. 20). It has been demonstrated that with the exception of core and lipid A determinants, all the other epitopes of smooth LPS are present in the NH (Aragó n et al., 1996b; Moreno et al., 1987). Therefore, it is not surprising that practically all infected animals with smooth Brucella strains develop similar levels of antibodies to NH and to the O-polysaccharide of the LPS; however, a proportion of the same animals do not develop detectable antibodies against core or lipid A determinants (Fig. 34). Despite this, the reaction of these molecules differs regarding the antibody-mediated complement activation. In sharp contrast with smooth LPS, the antibodies bound to NH are poor consumers of complement component C3b (E. Moreno and I. Moriyó n, unpublished observations). The failure of NH molecules to activate complement may be due to subtle differences in the degree of formylation between the smooth LPS Opolysaccharide and NH, rather than in the absence of core and lipid A, in the latter (M. Staaf et al., unpublished observations). Indeed, absorption assays performed to remove antibodies against core and lipid determinants showed that complement activation by smooth LPS could not be due to antibodies against these moieties. Abundant surface C3b receptors promote phagocytosis in activated macrophages (Wright and Silverstein, 1983). Reduced C3b opsonization could help to avoid ingestion by those cells in which brucellae would have a reduced chance of survival. Although NH mostly binds immunoglobulin G (IgG; Díaz-Aparicio et al., 1993), and macrophages bear Fcγ receptors, reduced C3b opsonization could be important at the onset of the infection when the predominant antibodies are IgM. At this time, Brucella could penetrate phagocytes by interaction of the smooth LPS and NH with the mannose-fucose receptor without triggering the production of toxic oxygen derivatives, as has been pointed out before (Campbell et al., 1994). This opsonin-independent route of entry could represent a strategy suitable for an intracellular pathogen because B. abortus cells either nonopsonized or opsonized with nonimmune serum fail to induce a significant oxidative burst in bovine polymorphonuclear leukocytes (Canning et al., 1988).

The Genus Brucella

B. melitensis

12

6

0 LPS

NH

R-LPS

Lipid A

Fig. 34. ELISA and radial immunodiffusion (RID) representing the IgG reactivity of B. abortus S19 vaccinated or B. abortus (biotype 1) naturally infected cows against smooth LPSs, rough LPSs, NHs and lipid as from B. abortus (biovar 1) and B. melitensis (biovar 1). Independent of the antigen attached (B. abortus or B. melitensis) to the ELISA plate or the NH used in the RID analysis, results are the same for both types of antigens. ELISA optical density values for B. abortus and B. melitensis antigens did not vary more than 5% in three different runs. RID was qualitatively evaluated as positive or negative. Notice that vaccinated animals do not react against NH, while infected animals do. Compare to (Fig. 19). The relevant epitopes in these reactions are the overlapping common epitopes in both B. abortus and B. melitensis antigens.

mally found in secretions, such as lysozyme, lactoferrin, lytic complement, chelating substances, proteolytic enzymes, phospholipase A2, defensins and defensin-like molecules, as well as to a variety of substances released by leukocytes during inflammation (Martínez de Tejada et al., 1995; Páramo et al., 1998; Riley and Robertson, 1984a). This resistance is related to the Brucella outer-membrane structure (mainly the LPS and

390

E. Moreno and I. Moriyó n

polysaccharide molecules), which from the chemical and functional point of view constitutes an excellent shelter for resisting the assault of these bactericidal mediators (Kreutzer and Robertson, 1979a; Martínez de Tejada et al., 1995). In addition to this natural resistance, Brucella cells or extracts obtained from these organisms do not significantly induce the exocytosis of neutrophil granules (Canning et al., 1986; Kreutzer et al., 1979b), suggesting functional modulation of these cells. Indeed, Brucella LPS does not stimulate the degranulation, respiratory burst or lysozyme release from neutrophils (Rasool et al., 1992). Moreover, GMPand adenine-containing Brucella extracts have been proposed to inhibit neutrophil degranulation (Canning et al., 1986). This last result, however, must be considered from the perspective that killed Brucella do not inhibit digestion by phagocytic cells. BRUCELLA Binds and Penetrates Host Cells Very few clear-cut studies have addressed the question of the mechanisms by which Brucella binds to and penetrates cells. Similarly to other pathogens (for review, see Méresse et al., 1999; Pizarro-Cerdá et al., 1997; Pizarro-Cerdá et al., 2000), Brucella organisms may have developed specialized microbial structures to gain access to host cytoplasmic compartments. In nonprofessional phagocytic epithelial cells, Brucella may induce its own uptake, since certain mutant bacteria are poorly internalized by the inoculated monolayers (Sola-Landa et al., 1998). Compared to other intracellular Proteobacteria, such as Salmonella, Shigella or Legionella, surprisingly low numbers of virulent Brucella bind to the surface of naive professional and nonprofessional phagocytes (Pizarro-Cerdá et al., 1998b; PizarroCerdá et al., 1998c; Sola-Landa et al., 1998). Despite this low attachment of virulent Brucella, their efficiency of penetration is close to 100%. As early as five minutes after infection, B. abortus is found within vacuoles of cells. Electron micrographs of Brucella penetrating cells have shown that phagocytosis can occur via a zipper mechanism (Ackermann et al., 1988; PizarroCerdá et al., 1999a; Fig. 33) or alternatively by a slow process resembling capping with little cell membrane rearrangements (Guzmán-Verri et al., 2001). Interaction with the Host Cell Membrane Sequences coding for putative proteins similar to those necessary to build flagella have been identified in the Brucella genome (DelVecchio et al., 2002; Sánchez et al., 2001; O’Callaghan et al., 1999), but none of these structures have ever been observed in Brucella cells, precluding

CHAPTER 3.1.16

their participation as attachment molecules. Alternatively, the involvement of nonfibrillar adhesins remains open, since sequences compatible with these protein molecules have been discovered in Brucella (DelVecchio et al., 2002; Ugalde, 1999; http://www.genome. scranton.edu.WIT2; www.tigr.org). Various studies have reported that nonvirulent smooth and rough Brucella mutants bind more readily and in larger amounts to the surface of professional and nonprofessional phagocytes than virulent strains do (Detilleux et al., 1990a; Detilleux et al., 1990b; Sola-Landa et al., 1998). However, there is not a strict direct correlation between the expression of O- and NH polysaccharides and binding to cells (Freer et al., 1999; Pizarro-Cerdá, 1998a). For instance, the virulent smooth Brucella and the “natural” rough B. ovis and B. canis (not expressing O-polysaccharide or NH) bind in very low number to HeLa cells. In contrast, the perosamine synthetase (perA) rough B. abortus mutants (not expressing Opolysaccharide or NH molecules) and the spontaneous rough Brucella mutants bind to these cells in much higher numbers than do smooth brucellae (Freer et al., 1999). Although difficult to interpret, the fact is that virulent Brucella cells have the tendency to attach in lower numbers to cultured epithelial cells than nonvirulent bacteria do (Fig. 21). The host cytoplasmic membrane directly adjacent to rough Brucella mutants is frequently thicker than the cell membrane adjacent to smooth bacteria and occasionally forms coatedpits (Detilleux et al., 1990a). These observations, although interesting, may not reflect the real physiological interactions between virulent smooth organisms and their host cells, since coated-pits adjacent to smooth Brucella have not been seen during phagocytosis. In contrast, increased adherence displayed by many mutant Brucella seems to be the result of altered outermembrane properties, which expose “new” sites for attaching in a nonphysiological manner (Allen et al., 1998; Corbeil et al., 1988; Detilleux et al., 1990a, 1990b; Freer et al., 1999). It is known that polysaccharide moieties in the outermembrane hide hydrophobic and ionic charges on the surface of the bacteria (Fig. 20) that obstruct membrane domains capable of participating in the nonspecific interactions between Brucella and the host cells (Aragó n et al., 1996b). In addition, the absence of surface Opolysaccharide and NH exposes core sugars to the external surface and gives more access to Omps, which may serve as ligand sites (Bowden et al., 1995; Cloeckaert et al., 1990). Invasion of Professional Phagocytes A scheme showing the binding, penetration and replication

CHAPTER 3.1.16

The Genus Brucella

391

MHC-II-rafts-LPS complexes M6PR MHC-II comparment

Phagolysosome

Cathepsin C3bReceptor LPS H+

Sec 61b calnexin

GCSF EEA1, CTB filipi, cd48 aerolysin

Action ER

Apoptosis inhibition

Lipid rafts cholesterol GM-1 Mann-Receptor LPS Fc-Receptor

Fibronectin Receptor

Fig. 35. Schematic model of B. abortus invasion and intracellular trafficking in macrophages. Brucella organisms bind to discrete sites via receptor molecules (FcR, C3bR, MannR, and fibronectin), some of which are unknown. Brucella penetrates by a zipper-like phagocytosis with moderate recruitment of actin filaments. The ingested bacterium initially follows its route (thick green arrows) to early phagocytic compartments (marked by EAA1) which may be acidified (H +) by acquisition of specific proton pumps (dotted arrows). Granulocyte colony-stimulating factor (G-CSF) is involved in promoting fusions between endocytic vacuoles and phagosomes (dotted arrows). In macrophages, most of the brucellae are routed (thick red arrows) to fuse with lysosomes (marked by cathepsin D and LAMP1) generating phagolysosomes where they are destroyed, whereas only a few bacteria follow the route (black thin arrows) to their replicating niche inside the endoplasmic reticulum (marked by Sec61β) following the autophagocytic (marked by Sec61β and LAMP1) pathway. The released LPSs merge with MHC-II compartments and the complexes MHC-II-LPS are transported to the cell membrane, inhibiting antigen presentation (crossed white arrow). Inhibition of apoptosis is induced in both infected and noninfected phagocytic cells (broken arrow). Steps indicating endoplasmic reticulum recruitment and Brucella replication within the endoplasmic reticulum have been adapted from unpublished results from Jean Celli of the CIML-Marseille-Luminy, France.

cycle of smooth Brucella in macrophages is presented in Fig. 35. Brucella cells bind to and penetrate more efficiently professional than nonprofessional phagocytes. Still, the number of Brucella organisms associated to the former cells is comparatively much lower (from one to four logs) than the numbers observed with other intracellular Proteobacteria, such as Salmonella, Shigella or Legionella. There are differences in the binding and internalization of Brucella by phagocytic cells (Pizarro-Cerdá, 1998a). Concomitantly, opsonized Brucella cells attach and are more readily ingested by phagocytes than are nonopsonized cells. This phenomenon is more clearly observed with activated than with nonactivated professional phagocytes (Arenas et al., 2000; Eze et al., 2000; Gross et al., 1998; Harmon et al., 1988; Young et al., 1985). As in the case of Mycobacterium (Schorey et al., 1997), it may be that highly opsonized Brucella (prozone effect) could take advantage of phagocytosis by Fc or complement receptors to invade cells expressing these receptors. However, opsonization seems to negatively affect the rate of survival and multi-

plication of Brucella within phagocytes, suggesting that Fc or complement receptor-mediated phagocytosis works in favor of the host cells rather than the bacteria (Caron et al., 1994b; Gross et al., 1998; Harmon et al., 1988; Harmon et al., 1989). Brucella LPS and NH do not activate the alternative complement pathway (Moreno et al., 1981; Hoffmann et al., 1984). Therefore, these polysaccharides do not serve as direct attachment of C3b on the bacterial surface. The fact that nonopsonized Brucella organisms bind, penetrate and reproduce in vivo and in vitro within phagocytes from nonimmunized animals (Campbell et al., 1994; Gross et al., 2000; Kuzumawati et al., 2000; Sola-Landa et al., 1998) indicates the existence of a receptor-ligand mechanism independent from the Fc and complement receptors. The exclusion of membrane negatively charged groups on the site of Brucella attachment in macrophages, followed by zipper-like phagocytosis, suggests specific interactions between both types of cells (Gay et al., 1981). Lipid rafts rich in cholesterol, glycosylphosphati-

392

E. Moreno and I. Moriyó n

dylinositol and GM1 gangliosides seem to provide a port of for Brucella entry into murine macrophages under non-opsonic conditions (Naroeni and Porte, 2002; Watarai et al., 2002). Brucella organisms (either opsonized or nonopsonized) are rapidly internalized by murine macrophages and human monocytes after inducing recruitment of actin filaments without generating marked structural changes but a general membrane ruffling (Gay et al., 1981; Kuzumawati et al., 2000; Watarai et al., 2002). Penetration of Brucella into bovine mononuclear phagocytes is inhibited by bacterial cell envelopes, antibody against the α chain of the MAC-1 integrin (CD11b), O-polysaccharide and denatured IgG, no matter whether the cells originated from cattle naturally resistant or susceptible to brucellosis. In addition, fibronectin, mannan and antibodies against C3 also inhibit the penetration of nonopsonized bacteria to phagocytes from brucellosis-resistant cattle (Campbell et al., 1994). Since the O-polysaccharide of LPS and mannan inhibit the binding of Brucella to bovine macrophages, it is likely that lectin-like receptors on the surface of phagocytic cells participate in the Brucella uptake. This idea also is supported by the observation that the strong binding of Brucella onto murine B lymphocytes is inhibited by α-methyl mannosamine and LPS (Lee et al., 1983). In addition, it has been demonstrated that phagocytosis of Brucella by macrophages is accompanied by secretion of fibronectin, which is then detected inside the phagocytic vacuole (Gay et al., 1986). Since fibronectin also blocks the invasion by Brucella, it is likely that this fibrous matrix protein could participate in the binding and penetration of Brucella in macrophages. Moreover, the intravacuolar fibronectin contributes to the electron-dense material that is commonly seen surrounding the phagocytosed brucellae, suggesting that this protein is coingested with the bacteria (Gay et al., 1981; Gay et al., 1986). Invasion of Non-Professional Phagocytes A model showing the penetration and intracellular cycle of Brucella within nonprofessional phagocytic HeLa cells is presented in Fig. 36. As expected, immune or native sera are not necessary for invasion of Brucella species into nonprofessional phagocytes (Anderson and Cheville, 1986a; Anderson and Cheville, 1986b; Anderson et al., 1986c; Detilleux et al., 1990a; Detilleux et al., 1990b; Pizarro-Cerdá et al., 1999b). In HeLa cells, B. abortus attaches to cellular extensions that are compatible with adhesion plaques and between cell-to-cell contacts and penetrates by a zipper-like mechanisms or by capping-like mechanisms with moderate rearrangement of the cell membrane (Guzmán-Verri et al., 2001).

CHAPTER 3.1.16

Some of these observations have been performed in HeLa cell monolayers previously intoxicated with C. difficile toxins B [TcdB] and BF [TcdBF] and then infected with Brucella. The TcdB-intoxicated cells retract their body, leaving cellular spikes attached to the substrate. In contrast, TcdBF induces cell rounding and retraction of cellular spikes from the substratum. Brucella organisms attach in normal numbers to the cellular spikes of TcdB-treated cells. On the contrary, in TcdBF-intoxicated cells, Brucella organisms bind only in reduced numbers, and when binding occurs, the bacteria locate on the cellular body. It has been proposed that α5β1 integrin mediates the adhesion of rough mutant B. abortus RB51 to bovine trophoblasts (Bress et al., 1996). However, this observation has not been reproduced in HeLa cells in spite of the highly adherent nature of mutant rough Brucella to cells and inert substrates (Pizarro-Cerdá, 1998a). Uptake of killed or alive Brucella by Vero cells is suppressed by inhibitors of energy metabolism (iodoacetate and dinitrophenol), inhibitors of receptor-mediated endocytosis (monodansylcadaverine, amantadine and methylamine) and by repressors of endosomal acidification (chloroquine, ammonium chloride and monensin). These drugs are capable of inhibiting penetration when added at the same time as the bacterial inoculum (8 h), but not when added after the inoculation period, suggesting that the infection process occurs via receptor molecules and requires energy input from the host cell (Detilleux et al., 1991). The participation of the cytoskeleton, second messengers and GTPases in the internalization of B. abortus to HeLa cells has been recently investigated (Guzmán-Verri et al., 2001). Phalloidin staining has revealed a modest recruitment of actin filaments in the site of Brucella attachment. Inhibition of actin filaments by drugs (Detilleux et al., 1991; Pizarro-Cerdá, 1998a; Guzmán-Verri et al., 2001) or by different clostridial toxins (TcdB, TcdBF, TcdA, and TcsLT) functionally modifying the actin cytoskeleton through interaction with small GTPases of the Rho family hampers internalization but not binding to cells (Guzmán-Verri et al., 2001). Infection also is inhibited by chemicals and toxins that increase the levels of cyclic-AMP (dibutyl-cyclic-AMP and Vibrio cholerae enterotoxin), but it is stimulated by toxins and chemicals that increase the levels of cyclic-GMP (Escherichia coli enterotoxin A and dibutylcyclic-GMP). This suggests an inverse relationship between these two second-messengers during Brucella infection. Similarly, wortmanin (which inhibits the PIP3 kinase) considerably reduces the internalization of Brucella by HeLa cells, suggesting involvement of PIP3-kinase

CHAPTER 3.1.16

The Genus Brucella Endoplasmic reticulum Sec61β, calnexin ribophorin

Late endosome Rab7, M6P-R

Autophagosome Sec61β, LAMP1

Cell cycle

PLC(?) Tyr-K MAPK PIP3-K Rho, Rac

?-R Virulent B. abortus

Cdc42 H+?

Actin

Phagocitosis

Early phagosome Rab5, EEA1

Phagolysosome cathepsin D LAMP1 Lysosome Cathepsin D LAMP1

393

CNF Cdc42 Rho, Rac

Ruffles

Fig. 36. Schematic model of B. abortus invasion and intracellular trafficking in epithelial HeLa cells. Brucella organisms bind to discrete sites of cells via unknown receptor molecules (???-R) and penetrate by a discrete phagocytosis with moderate recruitment of actin filaments, activation of small GTPases (Cdc42, Rac, and Rho), mainly Cdc42, and signals mediated by second messengers (Tyr-K, MAP-K, and PIP3-K). The ingested bacterium is initially routed (green arrows) to early phagocytic compartments (marked by Rab5 and EEA1) which may be acidified (H+) by acquisition of specific proton pumps (dotted arrows). In HeLa cells most of the ingested virulent brucellae are routed (thick red arrows) to the endoplasmic reticulum by the autophagocytic route (marked by LAMP1 and Sec61β), whereas only a few bacteria are directed (thin black arrows) to phagolysosomes. Some mutants (BvrS/BvrR) are defective in penetration (crossed arrow), others cannot avoid fusion of early phagosomes with lysosomes (BvrS/BvrR, VirB1-VirB10, and cgs), whereas others transit from autophagosomes to phagolysosomes (S19) or from early phagosomes to the cell membrane (nonpolar VirB10; blue thick arrow). Signals for apoptosis inhibition may be released from the Brucella replicating niche.

phosphorylation during this process (GuzmánVerri et al., 2001). Since the level of cyclic GMP usually increases when the inositol phospholipid pathway is activated, it is likely that binding of Brucella to cells also stimulates the generation of IP3 via phospholipase C activation. Other cellular kinases, such as tyrosine kinases and MAP kinases, seem to be required for physiological internalization to HeLa cells, since inhibition of these enzymes hampers bacterial penetration (Guzmán-Verri et al., 2001). Brucella abortus attaches in larger numbers and is internalized more efficiently after intoxication of HeLa cells with the cytotoxic necrotizing factor (CNF), which deamidates the small GTPases Rho, Rac, and Cdc42 and induces ruffles and stress fiber formation (Guzmán-Verri et al., 2001). In these cells, Brucella organisms bind to the ruffles as well as to discrete sites and penetrate through these structures. Moreover, HeLa cells transfected with constitutive negative mutant plasmids expressing defective Rho, Rac and Cdc42 proteins are considerably less infected than control cells. On the contrary, the positive counterparts of these small GTPases expressed in HeLa cells stimulate binding and penetration of Brucella. Virulent Brucella selectively stimulates the generation of Cdc42

GTPase, reaching its maximum at 30 minutes after bacterial contact with cells. This property may be related to an Omp of group 3, more likely Omp3a (Omp25) and/or Omp3b. This is supported by the fact that the nonvirulent B. abortus BvrS mutant does not stimulate any of the GTPases, in spite of its binding to cells (GuzmánVerri et al., 2001). Microtubule depolymerizing agents, such as nocodazole and colchicine, partially reduce the internalization but not the replication of Brucella in HeLa cells (Guzmán-Verri et al., 2001). This strategy for invading cells differs from those employed by other intracellular pathogens, such as Salmonella and Shigella. Despite the obvious differences between Brucella and Listeria, both types of pathogens seem to employ very similar strategies (Finlay and Cossart, 1997a). BRUCELLA Traffics and Replicates Within Host Cells Brucella organisms have been found to survive and replicate within membranebound compartments of professional and nonprofessional phagocytes (Fig. 30). Despite the tropism of these pathogens for reproductive organs, the bacteria also localize within cells of various tissues at later stages of the infection (Enright, 1990b). In vivo, B. abortus has been

394

E. Moreno and I. Moriyó n

described within bovine, caprine and murine trophoblasts, in caprine lymphocytes, M cells, and chicken embryo fibroblasts, as well as in a number of professional phagocytes lining different tissues (Ackermann et al., 1988; Anderson and Cheville, 1986a; Anderson et al., 1986c; Anderson et al., 1986b; Cheville et al., 1992; Cheville et al., 1996a; Detilleux et al., 1988; Holland and Pickett, 1956; Richardson and Holt, 1964). In vitro, B. abortus has been described to replicate within vacuoles of hamster kidney cells (Hatten and Sulkin, 1966a; Hatten and Sulkin, 1966b), primary cultures of bovine adult and fetal cells (Richardson and Holt, 1964), and a number of epithelial cells and macrophages (Baldwin and Winter, 1994; Caron et al., 1994b; Detilleux et al., 1990a, 1990b; Liautard et al., 1996; Pizarro-Cerdá et al., 1998b; Pizarro-Cerdá et al., 1998c; SolaLanda et al., 1998). Among the nonprofessional phagocytes, epithelial HeLa and fibroblastic Vero cell lines have been the most extensively used for studying Brucella replication and intracellular trafficking. Among professional phagocytes, murine J774, murine peritoneal macrophages, human monocytes, bovine mammary gland macrophages and human neutrophils have been the most widely used cells. M cells, neutrophils, nonactivated macrophages from newly infected hosts, activated macrophages from immune animals and nonprofessional phagocytes all serve a different purpose during the course of Brucella infection. While the translocation of ingested Brucella organisms occurs through M cells, the first line of defense against Brucella invasion is secured by neutrophils, short-lived cells with a strong capacity to phagocytize (Ackermann et al., 1988). Although neutrophils are not the preferred niche for Brucella replication, some of these intracellular bacteria are capable of withstanding destruction inside these leukocytes (Kreutzer et al., 1979b; Young et al., 1985). In turn, this event may favor the spreading of the parasite from neutrophils to other tissues (Ackermann et al., 1988; Enright, 1990a). In the second line of defense are the macrophages, which, like neutrophils, can destroy an important proportion of the ingested Brucella but may also serve as substrate for Brucella replication as well as vehicles for transportation to other tissues. In the pregnant animal, Brucella invades the erythrophagocytic trophoblasts, which are the preferred replicating host cells and the site from which the bacteria spread to the fetus (Anderson et al., 1986b; Anderson et al., 1986c; Tobias et al., 1993). Generally, immune individuals are capable of controlling the infection via stimulation of the macrophagic system. Depending upon the animal species, the humoral response may serve as an important aid for phagocytosis and for con-

CHAPTER 3.1.16

ducting the intracellular route of the phagocytosed bacteria to destructive compartments. Important findings revealing the interaction between Brucella parasites and their cells have been established. At the molecular level, the information is still incomplete and analysis of the bacterial factors and cellular receptors involved in the invasion process is necessary to understand the downstream steps observed during internalization and intracellular transit. Physical isolation and molecular examination of the different Brucella-containing compartments are necessary for the identification of the molecules that play a role in the original trafficking exhibited by this parasite. Survival Within Polymorphonuclear Neutrophils Neutrophils are the primary line of defense during Brucella invasion. These cells are capable of ingesting and killing Brucella faster and more efficiently than other cells are (Kreutzer and Robertson, 1979a; Riley and Robertson, 1984a; Riley and Robertson, 1984b). After incubation with neutrophils, virulent Brucella organisms are steadily destroyed. From 50 to 80% of the ingested bacteria are killed within 2 to 5 hours (Riley and Robertson, 1984a). This microbicidal activity seems to be more efficiently performed when bacteria are opsonized (Canning et al., 1988; Young et al., 1985). Once ingested, the bacteria are found within vacuoles, some of which already demonstrate fusions with lysosome-like granules (Figs. 30 and 33). At later times, fusions of azurophilic granules increase in most but not all Brucella-containing vacuoles. These events proceed without extensive neutrophil degranulation (Riley and Robertson, 1984b). Exudates from lesions, milk and blood from infected animals demonstrate Brucella within resident neutrophils with no signs of replication (Ackermann et al., 1988; Tobias et al., 1993). It seems that some intracellular bacteria are capable of resisting for several hours or even for days the microbicidal assault displayed by neutrophils, which in time may release live bacteria on sites where they could invade more temperate host cells (Ackermann et al., 1988). Life Within Non-professional Phagocytes The initial number of Brucella cells per infected epithelial Vero or HeLa cell is low, with one or two bacteria per cell (Detilleux et al., 1990a, 1990b; Pizarro-Cerdá et al., 1998c; Pizarro-Cerdá et al., 1998b; Fig. 21). Even if the bacterial inoculum is augmented in several logs, the rate of infection per cell remains low, suggesting that not all cells are permissive (Sola-Landa et al., 1998). Despite this, once the brucellae bind, the penetration efficiency is close to 100%. Immediately after phagocytosis, Brucella organisms localize within

CHAPTER 3.1.16

single membrane compartments generally containing only one bacterium. During the first hours (1–5), no clear signs of intracellular bacterial replication are demonstrated, and the rate of cellular infection remains relatively constant for the next 5–8 hours. After 24–48 hours, the number of infected cells may decrease down to one half, depending on the concentration of antibiotic in the culture media, suggesting that at early stages phagosomes containing Brucella may converge with readily pinocytosed vacuoles from the constitutive endocytic route. The number of bacteria, estimated as CFU, increases several logs from the initial inoculum, indicating extensive bacterial replication within the cytoplasm of infected cells. Although many cells possess a large number of intracellular bacteria, the cell monolayers do not show signs of cytopathic effect and the individual cells remain attached to the matrix. A small proportion of cells contain a low number of Brucella (1–50) with no signs of bacterial degradation, while fewer of them demonstrate bacterial debris within phagolysosomelike compartments. The small number of bacteria within cells is not the result of new infections, since normally low concentrations of gentamycin (0.5 µg/ml) are capable of controlling extracellular bacteria (Pizarro-Cerdá et al., 1998c; PizarroCerdá et al., 1998b). Alternatively, the low number of live bacteria within host cell vacuoles after 48 hours may be reminiscent of chronic infections in animals (C. Guzmán-Verri et al., unpublished observations). Multivesicular bodies compatible with autophagosomes are frequently present in infected cells but not in control monolayers. The absolute increase in CFU is due to an expansion in the number of intracellular bacteria per parasitized cell rather than to an increase in the number of infected cells (Detilleux et al., 1990a, 1990b; Pizarro-Cerdá et al., 1998b; Pizarro-Cerdá et al., 1998c). Under the electron microscope, the cell cytoplasm is filled with Brucella within ribosome-lined cisternae, compatible with the endoplasmic reticulum (Detilleux et al., 1990a, 1990b). In the most heavily infected cells, bacteria also are seen within the perinuclear space. Although the nuclear membrane is constrained due to the large number of bacteria surrounding the nucleus, none of them invade the cell nucleus. These heavily parasitized cells do not show degeneration, do not look apoptotic or present signs of necrosis (Chaves-Olarte et al., 2002). Moreover, dividing cells with the cytoplasm packed with bacteria are frequently observed (Fig. 30). Nevertheless, beyond 48 hours cellular rupture proceeds as a consequence of Brucella overgrowth. The antibiotics in the tissue culture medium kill the freed bacteria, generating a rapid decrease in the number of

The Genus Brucella

395

colony forming units (Pizarro-Cerdá et al., 1998b). Very similar events to those described in Vero and HeLa cells at 48 hours postinfection have been described in trophoblasts from experimentally infected animals (Anderson and Cheville, 1986a; Anderson et al., 1986b; Anderson et al., 1986c). Most if not all the Brucella cells are within the endoplasmic reticulum, and despite being filled with bacteria, infected trophoblasts do not show signs of degeneration, and the cellular junctions look normal (Fig. 30). Matching to what has been observed in cultured monolayers, bacteria remain within the endoplasmic reticulum compartments of individual cells, with no signs of horizontal movement from cell to cell. Actually, some noninfected cells are bound together with heavily infected trophoblasts, giving a clear indication that, even in vivo, Brucella organisms remain within vacuoles of their host cells until released. The trophoblast nucleus is never parasitized, although it is constrained by bacteria that lay very close or within cisternae that collide with the nuclear membrane. Similarly to epithelial cell monolayers, the infected trophoblasts seem to fracture at later times, owing to the large number of intracellular bacteria that eventually are released to the lumen. Transient Interaction with the Early Endosomal Network During the first minutes after invasion, both the virulent B. abortus strain 2308 and the attenuated strain 19 interact with an intracellular compartment related to the early endosomal network (Fig. 36). This is confirmed by the presence of markers, such as the transferrin receptor, the small GTP-binding protein rab5, or the early endosomal antigen 1 (EEA1), in the Brucellacontaining compartments (Pizarro-Cerdá et al., 1998). Several of these markers also have been observed in early phagosomes of other intracellular pathogens, such as Salmonella typhimurium, Leishmania donovani, Mycobacterium tuberculosis and Listeria monocytogenes (Alvárez-Domínguez and Stahl, 1999; SteeleMortimer et al., 1999; Scianimanico et al., 1999; Via et al., 1997). All these parasites use very different strategies to associate and penetrate host cells, suggesting that there is an invariable minimal cellular machinery regularly required to accomplish the internalization steps of an external agent. This seems to be more important when the host cell is the entity that performs an active role in the process. A different phenomenon could be expected in the case of the Brucella close relative Bartonella bacilliformis or the protozoan Toxoplasma gondii and Trypanosoma cruzi, in which the motile forces of the pathogens themselves drive the invasion processes

396

E. Moreno and I. Moriyó n

(Doborowski et al., 1996; Tardieux et al., 1992; Scherer et al., 1993). The association of B. abortus with the early endocytic network is transient, since after 10 min of internalization, the number of Brucellacontaining compartments labeled either with rab5 or EEA1 decreases significantly, and no labeling is detected with these markers after 30 min postinoculation (Pizarro-Cerdá, 1998a). The integrity of the early endosomal system is relevant to the subsequent normal trafficking of B. abortus in host cells. For instance, in the cell line NIH3T3 rab5Q79L, in which the activated form of rab5 (bound to GTP) is expressed, an important fraction of the internalized parasites are unable to escape from the early Brucellacontaining compartment supporting bacterial replication within giant vesicles labeled with rab5 (Pizarro-Cerdá, 1998a). However, after 48 h of infection, B. abortus proliferation is attenuated in NIH3T3 rab5Q79L cells in comparison to the wildtype NIH3T3 counterparts (PizarroCerdá, 1998a). The augmentation of the endocytic activity of the mutant cells could be responsible for an increase in the delivery of gentamycin to intracellular compartments. Indeed, an important fraction of the intracellular brucellae remains associated with the early endosomal network, exposing the bacteria to the bactericidal activity of the antibiotic within this compartment. Alternatively, this unnaturally generated compartment may not be suitable for the adequate delivery of the necessary nutrients for intracellular bacterial replication. Association with the Autophagic Machinery After their transit through early phagosomes, neither the virulent B. abortus strain 2308 nor the attenuated strain 19 interacts with the late endosomal network at 30 min postinvasion (PizarroCerdá et al., 1998b). In contrast, latex beads or dead bacteria-containing phagosomes interact transiently with late endocytic compartments, characterized by the presence of the small GTPbinding protein rab7 or the mannose 6-phosphate receptors. This result is confirmed after infection of NIH3T3 rab7Q67L cells (in which rab7 is expressed in its GTP-bound form) with B. abortus 2308. In this mutant cell line, the bacterial replication is similar to that in wildtype cells (Pizarro-Cerdá, 1998a). Although Brucella does not transit through the late endosome network, vacuole acidification seems to be required, since chloroquine, ammonium chloride and monensin (all substances that inhibit endosomal acidification) are capable of reducing the number of intracellular bacteria at early but not at later times after infection (Detilleux et al., 1991). Acidification step without the acquisition of lysosomal markers may be necessary for the activa-

CHAPTER 3.1.16

tion of virulence genes as it occurs with other parasites (Antoine et al., 1990; Buchmeier and Heffron, 1990). Following these initial steps, the Brucellacontaining compartment is transformed gradually, and after 1 h of internalization, both virulent strain 2308 and attenuated strain 19 are present in an intracellular multimembranous compartment decorated with the lysosomal-associated membrane protein (LAMP) 1, but devoid of the luminal lysosomal hydrolase cathepsin D. This finding supports previous propositions in the sense that virulent B. abortus inhibits the fusion of its phagosome with lysosomal compartments (Frenchick et al., 1985). Several criteria permitted the identification of this late Brucellacontaining compartment as an autophagosome (Fig. 36). First, the multimembranous nature of the LAMP-1-positive cathepsin D-negative vacuole is highly reminiscent of autophagosomes. Second, this compartment is labeled by monodansylcadaverine, a marker known to accumulate in autophagosomal bodies. Third, the endoplasmic reticulum marker sec61β is present in this Brucella-containing vacuole, attesting to an endoplasmic reticulum-related origin of this compartment. Fourth, modulation of the autophagocytic process regulates the intracellular fate of the internalized brucellae (Pizarro-Cerdá et al., 1998b; Pizarro-Cerdá et al., 1998c). The presence of LAMP-1 in the late Brucellacontaining compartment could be explained by a direct delivery of this molecule from the Golgi complex to the maturing autophagosomes. It is interesting to note that LAMP-1 has been widely associated with pathogen-containing compartments, such as the vacuoles of S. typhimurium, L. donovani and L. monocytogenes (Scianimanico et al., 1999; Steele-Mortimer et al., 1999; J. Pizarro-Cerdá, personal communication). This molecule could be present in these compartments accidentally as a bystander, only as an outcome of the different trafficking pathways followed by this molecule, transported in certain cases to the plasma membrane before being delivered to the lysosomes (Hunziker and Geuze, 1995). However, the actual function of the LAMP family of glycoproteins has not been clearly defined (Andrejewski et al., 1999), and it would be interesting to determine if this molecule actually plays an active role that could be relevant to the intracellular survival of all the intracellular pathogens described above. The association of an intracellular pathogen with the autophagic pathway is not unique to B. abortus, and it has also been observed in the case of Legionella (Swanson and Isberg, 1995). It is not known how these bacteria are able to interact with the autophagic cascade. An interaction between early endocytic compartments and

CHAPTER 3.1.16

autophagic vacuoles has already been detected (Liou et al., 1997), indicating that a physical connection could exist between early Brucellacontaining compartments and autophagosomes. Several scenarios could then be conceived to explain the transfer of the pathogen from one intracellular compartment to the other. First, there could be a fusion between the Brucellacontaining compartment and an already formed autophagic vacuole. However, how the bacteria are finally found within the luminal space of a multimembranous compartment could not be directly explained by this hypothesis. A second, but highly improbable, possibility is the escape of B. abortus from the Brucella-containing compartment to the cytoplasmic space, where the bacteria could be captured by nascent autophagosomes, but free brucellae are seldom observed in the cytosol of infected cells. A third possibility would be that the autophagosomal vacuoles are formed by invagination of the endoplasmic reticulum membranes around Brucella-containing compartments. Nevertheless, the absence of other endoplasmic reticulum markers, such as BiP or ribophorin, in the Brucella-containing compartment (Pizarro-Cerdá et al., 1998b) contradicts this hypothesis. A modification of this alternative would be that only specialized regions of the endoplasmic reticulum, devoid of BiP or ribophorin, are involved in autophagosome formation. Replication Within the Endoplasmic Reticulum In

contrast to what is observed with attenuated and killed bacteria, most of the intracellular virulent Brucella-containing phagosomes lose the LAMP-1 labeling and never acquire lysosomal markers in nonprofessional phagocytes (PizarroCerdá et al., 1998b; Pizarro-Cerdá et al., 1998c). However, this final Brucella-containing compartment retains the sec61β labeling and acquires other markers of the endoplasmic reticulum, such as the protein disulfide isomerase and calnexin (Pizarro-Cerdá et al., 1998b). The morphology of the final Brucella-containing compartment also differs from that of the autophagosomal stage: only a single membrane is detected around the replicating brucellae, and their intracellular location corresponds to the perinuclear area of the infected cells (PizarroCerdá et al., 1998b; Pizarro-Cerdá et al., 1998c). All these data suggest that the virulent B. abortus transits from autophagosomes to the endoplasmic reticulum of host cells, where actual bacterial multiplication occurs (Fig. 36), confirming previous ultrastructural studies in trophoblasts and other mammalian cell lines (Anderson and Cheville, 1986a; Anderson and Cheville, 1986b; Detilleux et al., 1990a, 1990b). Additional evidence confirms the nature of the final niche of B.

The Genus Brucella

397

abortus replication in host cells. First, treatment of infected cells with brefeldin A, which normally induces the reorganization of the Golgi complex around the endoplasmic reticulum, induces the colocalization of Golgi markers around the Brucella-containing compartments (Pizarro-Cerdá et al., 1998b). Second, the treatment of infected cells with proaerolysin, a drug from Aeromonas hydrophyla that induces vacuolization of the endoplasmic reticulum in target cells, induces vacuolation of Brucella-containing compartment (Pizarro-Cerdá et al., 1998b). Treatment of target cells with proaerolysin before Brucella inoculation impairs the bacterial replication process and induces the degradation of virulent strain 2308 (Pizarro-Cerdá et al., 1998c), suggesting that the integrity of the structure of the endoplasmic reticulum is indispensable for the appropriate multiplication of B. abortus. The benefits involved in the association of intracellular pathogens with the host-cell endoplasmic reticulum have not been characterized yet. Besides B. abortus, L. pneumophila, T. gondii and simian virus 40 multiply in this intracellular environment, revealing a path of convergent evolution in nonrelated organisms. In addition to being a strategy for avoiding lysosomal fusion during the final steps of intracellular invasion, association of B. abortus with the host endoplasmic reticulum could be a means of obtaining metabolites synthesized or translocated to this compartment (Sinai et al., 1997a; Stang et al., 1997; Swanson and Isberg, 1995). The strategy would be to take advantage of the biosynthetic enzymes, protein-conducting channels or peptide pores to increase the local nutrient supply (Sinai, 1997b), fulfilling the nutritional requirements for the bacterial growth. A possible prediction of this model would be that the cellular stock of short-lived molecules would decrease in B. abortus-infected cells due to the shortage of amino acids and peptides or to the blocking of the biosynthetic process of new proteins in the endoplasmic reticulum. However, heavily infected cells show no decrease in stock levels of short-lived molecules, such as LAMP-1 and LAMP-2 (J. PizarroCerdá, personal communication). As with other intracellular parasites, such as S. typhimurium or Leishmania, it is still not clear why certain intracellular locations are preferred by different subsets of intracellular parasites to proliferate within host targets. Trophoblasts as a Source of Growth Factors In the

pregnant animal, Brucella organisms preferentially replicate in placental trophoblasts during the middle and late stages of gestation (Fig. 32), only after these cells actively secret steroids. The reason for this affinity and the process leading to

398

E. Moreno and I. Moriyó n

abortion after midgestation are not known. Infected trophoblasts produce cortisol, a steroidal hormone not normally generated by the placenta (Enright and Samartino, 1994). The increased levels of prostaglandin F2α and decreased production of progesterone, coupled with increased synthesis of estrogens and cortisol in the B. abortus-infected trophoblast at midand late-stages of gestation, are identical to the hormonal changes occurring at term in normal cattle with the initiation of parturition. Intracellular Brucella probably induces synthesis of steroids and modifies the metabolism of prostaglandin precursors, such as arachidonic acid, because these hormones may be used as growth factors by the bacteria. The increased hydrophobicity of the Brucella outer-membrane (Fig. 22), together with the preference of the bacteria for replicating within this cistern (Anderson et al., 1986b; Anderson et al., 1986c), may represent an evolutionary adaptation for using hydrophobic substances available within the endoplasmic reticulum of trophoblasts. Life Within Macrophages Taking apart the innate differences in susceptibility among the various species and strains of mammals, as well as the different sources from which macrophages could be isolated, it is important to understand that intracellular trafficking and replication of Brucella within these cells could be studied from various dichotomous perspectives. Each of these dichotomies possess their own variables; for example, activated versus resting macrophages, immune versus naïve macrophages, phagocytosis of opsonized versus nonopsonized bacteria, and cell cultures in the presence or in the absence of antibiotics. It is obvious that combined models of these dichotomies are possible, complicating matters even more. Therefore, it is important to understand that each of the various scenarios may alter the intracellular trafficking and the replication of the ingested bacteria (Eze et al., 2000; Jiang and Baldwin, 1993a; Jiang et al., 1993c). Similarly to the ingestion of Brucella, the microbicidal activity seems to be better performed with opsonized rather than with nonopsonized bacteria and by activated rather than by nonactivated macrophages (Pomales-Lebron and Stinebring, 1957). In addition, the presence of antibiotics considerably alters the outcome of the infection, depending upon their concentration in the culture medium (Eze et al., 2000). In this sense, it is expected that bacterial opsonization and macrophage activation determine, to some extent, the outcome in the intracellular trafficking and the fusion events of vacuoles containing Brucella. With respect to opsonization, caution is necessary, since conflicting results have been obtained.

CHAPTER 3.1.16

Structural and Morphological Changes During Macrophage Infection During the first two hours after macrophage Brucella infection, the bacteria localize within single phagosomes, with no signs of microbial degradation (Arenas et al., 2000; Pizarro-Cerdá, 1998a; Pizarro-Cerdá et al., 1998b; Pizarro-Cerdá et al., 1999b). The number of ingested Brucella depends upon the inoculum size, opsonization and macrophage activation (Kuzumawati et al., 2000). Fusions between lysosome-like granules and some Brucellacontaining phagosomes are evident at these early times, and few intracellular bacteria localize within multimembrane compartments. After 12–15 h of ingestion, the number of Brucella decreases one to two logs, and more than one bacterium per vacuole is frequent (Gross et al., 1998; Gross et al., 2000; Jiang and Baldwin, 1993a; Jiang et al., 1993c; Porte et al., 1999). Microscopic observation reveals that an important proportion of Brucella-containing compartments have fused with lysosomal granules and many of the bacteria within look degraded, although a few appear intact, generally surrounded by a single vacuolar membrane (Arenas et al., 2000; Gay et al., 1981; Gay et al., 1986; Harmon et al., 1988). At later times (15–24 h), the number of intracellular bacteria increases in some but not all infected cells. While some macrophages are capable of controlling the infection, others become an adequate substrate for Brucella replication (Gay et al., 1981; Gay et al., 1986; Pomales-Lebron et al., 1957). Still, some bacteria look digested within vacuoles and bacterial debris are evident, but a proportion of nondegraded bacteria remain, generally surrounded by a single vacuolar membrane. Between 24 and 48 hours, the number of intracellular Brucella per cell rises, until the cytoplasm of the phagocytic cells is filled with parasitic bacteria (Jiang and Baldwin, 1993a; PizarroCerdá, 1998a; Pizarro-Cerdá et al., 1999a; Pomales-Lebron et al., 1957). Similar to what has been described for nonprofessional phagocytes, the nuclear membrane looks constrained, owing to the large number of microorganisms surrounding the nucleus. However, none of the bacteria invade the cell nucleus (Fig. 30). These heavily parasitized macrophages do not look apoptotic, look vacuolated or present signs of necrosis. Nevertheless, beyond 48 h, cellular rupture seems to proceed as a consequence of Brucella overgrowth, and release of the bacteria occurs. The freed bacteria are normally killed by the antibiotics in the tissue culture medium, generating a rapid decrease in the number of colony forming units (Sola-Landa et al., 1998). Trafficking and Replication Within Macrophages The fact that not all cultured infected macrophages sustain bacterial replica-

CHAPTER 3.1.16

tion after 12 h is an indication of the powerful brucellacidal activity of these cells. Those bacteria that finally replicate intracellularly must escape from the digestive abilities of macrophages. It has been proposed that phagosome acidification dependent of proton ATPases, occurring at early (1 h) but not at later times (after 7 h), is essential for bacterial replication (Porte et al., 1999). Other investigators have observed that acidification of phagosomes takes place at early (0.5–2 h) as well as later (20 h) times (Arenas et al., 2000). Acidification of Brucella-containing phagosomes may work in two directions. One of them may just be the default outcome of phagosome-lysosome fusion with negative consequences for the bacterial survival. The second may work in favor of the parasitic bacteria. That is, the intracellular bacterium may require an acidic environment for activating genes necessary for an intracellular life cycle, such as the type IV secretion system apparatus (Alpuche-Aranda et al., 1992; Boschiroli et al., 2002). Even though lowering the pH of the phagosome may be necessary to prepare the Brucella replicating environment, this may not be sufficient, since phagosomes containing live bacteria or killed organisms acidify at a similar rate. Another consideration is the presence of gentamycin in the culture media. This antibiotic is readily pinocytosed by macrophages and considerably inhibits the intracellular growth of the Brucella, mainly when it is used in the culture medium at concentrations above 10 µg/ml (Eze et al., 2000). The biogenesis of phagolysosomes and the ability to degrade invading microorganisms involve a regulated series of interactions between phagosomes and endocytic organelles (Desjardins et al., 1994; Finlay and Falkow, 1997b; Pitt et al., 1992). During the first stages after infection (0.5–2 h), an important proportion of phagosomes containing bacteria seem reluctant to fuse with newly internalized vesicles, with the characteristics of lysosomes (PizarroCerdá et al., 1999a; Pizarro-Cerdá, 1998a). At ten minutes after inoculation, Brucella organisms are transiently detected in phagosomes, characterized by the presence of EEA1 (Fig. 35). Live but not dead Brucella exclude annexin I from early macrophage phagosomes (Harricane et al., 1996). At one hour postinoculation, bacteria are located within a compartment positive for LAMP but negative for mannose-6-phosphate receptor and cathepsin D protein, indicating that virulent Brucella avoids fusion with late endosomes and lysosomes (Pizarro-Cerdá et al., 1999a; Pizarro-Cerdá, 1998a). At later times (12– 24 h), fusions between newly internalized vesicles and Brucella-containing vacuoles are common events. However, the lysosomal mark-

The Genus Brucella

399

ers LAMP and cathepsin D are excluded from vacuoles containing replicating Brucella (Pizarro-Cerdá, 1998a), indicating that some bacteria have actively avoided the constitutive degradative pathway commonly followed by inert particles (Desjardins et al., 1994). Mature compartments containing live replicating Brucella (after 24 h) are devoid of the late endosomal marker mannose-6-phosphate receptor and lysosomal proteins (LAMP and cathepsin D) but are positive for the endoplasmic reticulum marker sec61β (Fig. 35), suggesting that at least some bacteria have reached the same replicating niche (the endoplasmic reticulum) as in nonprofessional phagocytes (Pizarro-Cerdá et al., 1999a; Pizarro-Cerdá, 1998a). In general terms, phagocytic cells are more prone to destroy Brucella and control its intracellular replication than are nonprofessional phagocytes. For instance, when Brucella strains of low and high virulence are compared, it is clear that no strain displaying low virulence withstands the powerful destructive machinery of professional phagocytes, whereas some of these strains are capable of replicating within nonprofessional phagocytic cells (Detilleux et al., 1990a; Detilleux et al., 1990b; Harmon et al., 1988; Fig. 35). Similarly, macrophages defective in fusion events are more permissive cells not only for virulent Brucella, but also for the attenuated strains that are killed by healthy macrophages (Pizarro-Cerdá et al., 1999b). Moreover, microscopic examination reveals that while in nonprofessional phagocytes most of the internalized bacteria travel within intracellular compartments that do not fuse with late endosomes and lysosomes, in macrophages a relatively large proportion of the phagosomes fuse with acidic vacuoles and lysosome-like compartments (Arenas et al., 2000). In nonprofessional phagocytes, most of the ingested Brucella transit from early endosomes to autophagosomes and then to the endoplasmic reticulum (Fig. 35), whereas in macrophages just a small proportion of Brucella-containing compartments seem to reach the endoplasmic reticulum (Fig. 35). In murine knockout macrophages deficient in fusion events, both virulent and attenuated B. abortus strain 19 replicate more readily than in normal macrophages (Pizarro-Cerdá et al., 1999a). Restoration of this defect re-establishes the microbicidal activity, indicating that fusion of vacuoles containing Brucella with newly internalized vesicles is necessary for controlling intracellular Brucella replication. Therefore, it seems that Brucella within nonfusiogenic vacuoles are more likely to reach the final intracellular replicating niche than those bacteria within vacuoles that fuse with lysosome-like compartments.

400

E. Moreno and I. Moriyó n

Electron microscopy of infected macrophages reveals that the replicating pattern displayed by Brucella within macrophages, at later times (Fig. 30), is reminiscent of the replicating pattern within the endoplasmic reticulum of trophoblasts and epithelial cells (Detilleux et al., 1990b; Anderson and Cheville, 1986a; Anderson and Cheville, 1986b). Some investigators have proposed that in macrophages, Brucella organisms do not reach the endoplasmic reticulum but rather they delay maturation of phagosomes into phagolysosomes and that, during this process, acidification and subsequent modification of the compartment establish the replicating niche, which is compatible with a modified phagolysosome (Arenas et al., 2000). Others have proposed that virulent Brucella transit from early phagosomes to the endoplasmic reticulum in both macrophages and nonprofessional phagocytes (Pizarro-Cerdá, 1998a; Pizarro-Cerdá et al., 1999b). Although at this point these two views cannot be conciliated, it is clear from the pure microscopic point of view that a parallelism in the replication of Brucella within the cytoplasm of macrophages and nonprofessional phagocytes exists at later times. It may be that most of the Brucella organisms in macrophages are destroyed by lysosomes, while only a few bacteria arrive to the endoplasmic reticulum. However, once these few bacteria are within the endoplasmic reticulum, they are capable of replicating inside this compartment, which is nonfusiogenic with bactericidal lysosomes (Fig. 35). In nonprofessional phagocytes, the events are reversed: only a few internalized Brucella are destroyed within lysosomes, while most of the Brucella arrive to the endoplasmic reticulum, trafficking through the safer autophagocytic route (Fig. 36). Obviously, the most economical pathway for the bacteria would be to use the same molecular machinery for intracellular trafficking and for subtracting resources in different host cells, rather than to possess a different molecular strategy for each cell type. An example of this can be seen in Legionella species, bacteria that use the same machinery and strategy to parasitize their free-living host amoebae and the resident macrophages in the lung of their victims (Barker et al., 1993). Thinking in this direction, the phagolysosome route establishes different constraints that must be surpassed. Similarly, the phagolysosomes and endoplasmic reticulum perform different functions and display a different set of resources for the intracellular bacteria. Therefore, it seems unlikely that Brucella could have evolved two different mechanisms to adapt equally well to two distinct intracellular environments as proposed by some authors (Arenas et al., 2000).

CHAPTER 3.1.16

Cellular Functions During Infection Transit of virulent Brucella from autophagosomes to the endoplasmic reticulum requires cellular machinery that has not been identified. In CNFintoxicated cells, the intracellular trafficking to the endoplasmic reticulum via the autophagocytic route remains intact, even though the actin cytoskeleton is functionally arrested (C. Guzmán-Verri et al., unpublished observations). Similarly, segregation of chromosomes and nuclei assembly are not defective, whereas the formation of a contractile ring for cell cytokinesis, mediated by an actin-myosin structure, is impaired in these CNF-intoxicated cells. Colchicine added during the infecting period has a modest effect on internalization, although it reduces the number of cells with intracellular replicating bacteria and has a profound effect on the morphology of the parasitized cells, generating polymicronuclei (Detilleux et al., 1991). It is possible that microtubule but not actin structures are involved in the retrograde transport of Brucella-containing compartment to the endoplasmic reticulum. The participation of molecular motors, such as dynein, which normally promotes the retrograde motion of membranebound vesicles through microtubules, is a mechanism that must be considered during the intracellular biogenesis of Brucella. Cycloheximide does not inhibit intracellular bacterial replication, and therefore, it is feasible to propose that de novo host protein synthesis is not required during Brucella parasitism (Detilleux et al., 1991). Proteins from the coat protein I (COPI) complex, known to participate in the anterograde transport of vesicles in the Golgi apparatus, have recently been implicated in the retrograde transport of vesicles to the endoplasmic reticulum (Orci et al., 1986; Orci et al., 1997). The presence of several subunits of the COPI complex in compartments associated with the endocytic cascade (Whitney et al., 1995) suggests that these molecules could establish a link among endocytic compartments, autophagosomal vacuoles and the endoplasmic reticulum, a link that is necessary for the intracellular trafficking of B. abortus. Molecules from the SNARE family (SNAREs are membraneassociated proteins that play a central role in vesicle targeting and intracellular membrane fusion in eukaryotic cells) or small GTP-binding proteins of the rab family could also be implicated in the retrograde transport of the Brucella-containing compartments. Recently, it has been demonstrated that autophagocytosis could be blocked by GTP-γS (a nonhydrolyzable analogue of GTP), suggesting that this process requires GTP-binding proteins (Kadowaki et al., 1994). It is interesting to note that the

CHAPTER 3.1.16

anterograde transport of vesicles from the endoplasmic reticulum is dependent on the small GTP-binding protein Sarpl and the COPI-like complex, COPII (D’Enfert et al., 1991). Similar or the same molecules could be implicated in autophagosomal formation and could be under B. abortus control to induce its retrograde transport to the endoplasmic reticulum. BRUCELLA Virulence Mechanisms Virulence can be understood as those bacterial mechanisms that cause cell damage or resist host defenses. Within this context, virulence factors are envisioned as bacterial products, structures or genes that, when absent or modified, alter the course of the pathogenesis without affecting the bacterial viability or the growth under regular culture conditions. There are overwhelming data demonstrating that the ability of Brucella organisms to produce disease relies in their capacity to invade cells of healthy immunocompetent hosts (Baldwin and Winter, 1994; Enright, 1990b; Liautard et al., 1996; Pizarro-Cerdá et al., 1999b; Pizarro-Cerdá et al., 2000). Two simple conclusions are derived from these clear cut facts: first, Brucella organisms are primary pathogens of mammals that do not require anomalous host conditions to establish infections; and second, the essential Brucella virulence strategy can be envisioned as the ability to replicate within host cells until more virulent bacteria are released. Therefore, the maintenance of these properties may be considered sine qua non for Brucella virulence and for successful parasitism. In spite of the relatively large number of DNA sequences identified as potential virulence genes, very few structures and mechanisms related to pathogenesis have been described (Moreno and Moriyó n, 2002; Sánchez et al., 2001; Ugalde et al., 1999). The Brucella mutants may be distinguished according to their replicating abilities in bacteriological media, cells and animals. Many of the attenuated mutants in cells or animals are metabolically defective strains displaying anomalous growth in vitro and requiring particular supplemented media and culture parameters. Others are impeded from making proteins necessary for confronting growth under stress conditions, whereas others have defects in repairing systems and division machinery. Although these general defects are interesting and prime for practical purposes, they are unlikely to be considered as virulence factors. Paragraphs in the sections Mechanisms of Entry to Host Cells through Modulation of the Immune Response summarize what it is known about the strategies followed by pathogenic intracellular Brucella and some of the genetic systems involved.

The Genus Brucella

401

Mechanisms of Entry to Host Cells Only efficiency of invasion and intracellular replication, but not adherence to the cell surface, positively correlate to Brucella virulence (Fig. 21). As stated (Antitumoral and Antiviral Activity of Brucella Cells and Their Fractions), a number of natural or artificial engineered nonvirulent rough bacteria bind and penetrate cells in a proportion larger than that of wildtype Brucella (Detilleux et al., 1990a; Detielleux et al., 1990b; Freer et al., 1999; Pizarro-Cerdá, 1998a; Sola-Landa et al., 1998). Mutations in bvrS or bvrR genes (The BvrR-BvrS System) of a two-component, sensory-regulatory system (Fig. 28) hamper the penetration of smooth B. abortus into HeLa cells and considerably reduce the efficient invasion of murine macrophages without impeding bacterial binding (Sola-Landa et al., 1998). Dysfunction of either the bvrR or bvrS impairs the bacterial invasion, intracellular trafficking and virulence, while knockout of both bvr genes seems to be lethal (D. O’Callaghan, personal communication). Both of these avirulent smooth mutants have the tendency to accumulate on the surface of cells. The rough perA mutant (Table 5), deficient only in the expression of surface O-polysaccharide and NH (J. Pizarro-Cerdá et al., unpublished observations), binds to cells in larger numbers. However, the double rough/bvrS mutant binds to cells in the same proportion as the bvrS single mutant (C. Guzmán-Verri et al., unpublished observations). Therefore, the exposed elements on the surface of rough Brucella necessary for adherence to cells are absent in the bvrS mutant. Comparative studies reveal that bvrS/bvrR mutants are deficient in at least two sets of Omps (C. Guzmán-Verri et al., unpublished observations), which correspond to the group 3 of Omps present in outer-membrane blebs (Gamazo et al., 1989; Moriyó n et al., 1987; Omp3a and Omp3b Family). Estimation of the Omp3a (Omp25) and Omp3b mRNA reveals low expression in both mutants, suggesting that the absence of this protein from the outer-membrane may be the result of reduced transcription. The role of Omp3a in virulence is further suggested by the fact that disruption of the omp3a gene attenuates Brucella (Elzer et al., 2000). Functional and structural analysis of the bvrS and bvrR mutants indicate that the lipid A and maybe the core, but not the O-polysaccharide of the LPS, have subtle but detectable changes with respect to the wildtype molecule. In this respect, the function of the BrvR-BrvS two-component regulatory system may be similar to that of other regulatory systems described in Gram-negative intracellular bacteria (Gunn and Miller, 1996). Although the BvrRBvrS has low homology with PhoP-PhoQ, it is interesting that, among other characteristics, the PhoP-PhoQ activates the PmrA-PmrB system,

402

E. Moreno and I. Moriyó n

which in turn, regulates the lipid A structure. The overall conclusion is that multiple genes, several of them important for the expression of Omps and LPS synthesis, may be under the control of bvrR-bvrS, and that these elements are important for virulence. It has been proposed that the type IV secretion system coded by the virB genes is involved in the penetration of Brucella to murine macrophages (Watarai et al., 2002); however, other investigators have indicated that the virB operon is not involved in cell invasion but rather in intracellular trafficking (Boschiroli et al., 2002; Comerci et al., 2001; Delrue et al., 2001; Sieira et al., 2000). Controlling the Host Cell Eukaryotic cells have evolved strategies to destroy microorganisms within phagolysosomes by direct action of microbicidal substances. Intracellular survival requires the invader to be capable of escaping and resisting this cellular machinery. According to their functional roles, neutrophils and activated macrophages are the primary and ultimate defense cells against Brucella infections, respectively. Depending upon the initial inoculum, the bacterial strain and the host animal, it is likely that an ample number of Brucella infections are controlled on site by neutrophils (Fig. 33). In these cells, the engulfed Brucella seldom divides, but it must resist the assault of intracellular microbicidal molecules to survive. In the nonimmune host, Brucella replicates within näive macrophages and nonprofessional phagocytic cells. Once the infection has been established, the activated macrophages, through a specific immune response, are the cardinal cells responsible for controlling and killing the invading Brucella. It seems that the difference in behavior observed among the various Brucella-infected cells is just a matter of degree in their killing capabilities. The survival strategy employed by Brucella within various cells seems to be basically the same and it can be resumed in the different but connected major episodes. Brucella Resists Killing Mechanisms Professional phagocytes, such as polymorphonuclear leukocytes and macrophages, are adapted to engulf and destroy bacteria within phagolysosomes by direct action of bactericidal mechanisms that involve a collection of enzymes and cationic peptides and production of reactive oxygen and nitrogen intermediates. The respiratory burst involves a series of enzymatic reactions responsible for the conversion of oxygen to active metabolites, some of which exhibit microbicidal activities. It has been observed that ingestion of nonopsonized B. abortus by phagocytes induces very little respiratory burst and a very poor production of reactive nitrogen intermediates

CHAPTER 3.1.16

(Caron et al., 1994b; Jiang and Baldwin, 1993a; Jiang et al., 1993c; Kreutzer et al., 1979b). Alternatively, opsonized B. abortus organisms exposed to activated phagocytic cells stimulate significant levels of both superoxide-ion production and myeloperoxidase-hydrogen peroxide halide activity (Harmon et al., 1987; Harmon et al., 1988; Jiang and Baldwin, 1993a; Jiang et al., 1993c; Young et al., 1985). Lysosomal proteins and bactericidal cationic peptides fail to kill Brucella organisms (Kreutzer and Robertson, 1979a; Kreutzer et al., 1979b; Martínez de Tejada et al., 1995). Even attenuated Brucella are more resistant to the action of these molecules than are other intracellular parasitic Proteobacteria, such as polymyxin B resistant Salmonella, indicating by this the evolutionary adaptation of Brucella to intracellular life. Outer Membrane Versus Bactericidal Substances The

most conspicuous structural defect that renders Brucella organisms avirulent is the absence of the O-polysaccharide and the concomitant absence of the related NH polysaccharide molecules. In other words, a defect that results from the dissociation from smooth to rough phenotype (Allen et al., 1998; Detilleux et al., 1990a, 1990b; Freer et al., 1996; Kreutzer et al., 1979b; Martínez de Tejada et al., 1995; Riley and Robertson, 1984a; Stevens et al., 1994a). In general, smooth Brucella are more resistant than rough strains to the killing action of bactericidal substances of phagocytes. The role of LPS in the permeability properties and in resistance to bactericidal substances has been definitively established in a series of experiments involving Brucella strains and the construction of LPS chimeras (Freer et al., 1996; Martínez de Tejada et al., 1995; Páramo et al., 1998). When the heterologous LPS inserted in the outer-membrane of susceptible bacteria corresponds to the less sensitive smooth B. abortus, the chimeras are more resistant to bactericidal cationic molecules. In contrast, when LPS is from the more sensitive bacteria, the chimeras are more susceptible to the action of bactericidal peptides. There is a direct correlation between the amount of heterologous smooth LPS on the surface of chimeric cells and sensitivity to bactericidal substances (Freer et al., 1996). Although this particular resistance to bactericidal molecules is related to the core and lipid A structures, there is a contribution of the O-polysaccharide and the associated NH, as suggested by the difference in susceptibility between the rough and smooth Brucella strains (Freer et al., 1996; Martínez de Tejada et al., 1995). It is worth noting that the resistance of Brucella LPS to cationic molecules is more conspicuous when it is integrated in its native outer-membrane. This is dem-

CHAPTER 3.1.16

onstrated by the fact that Brucella smooth LPS micelles are partially permeabilized by the action of bactericidal peptides, whereas Brucella cells are not (Freer et al., 1996). The O-polysaccharide and the NHs are only two of several factors necessary for virulence. Dysfunction of the BrvR-BrvS two-component, sensory regulatory system generates outermembrane alterations manifested as increased susceptibility to bactericidal cationic substances and surfactants (Properties of the Outer Membrane). These defects are partially restored by inserting wildtype LPS in the outer-membrane of the bvr mutants (E. Moreno et al., unpublished observations), reinforcing the idea that, in addition to LPS, other outer-membrane molecules are also important. Because bactericidal cationic peptides are molecules devoted to cell defense against pathogenic parasites (Vaara, 1992), it is likely that some of the outer-membrane features relevant for resistance to lysosomal substances are under the control of the BvrR-BvrS system. Stress Responses Brucella organisms generate a

collection of proteins that theoretically could inhibit the bactericidal action induced during the respiratory burst, as well as enzymes that could neutralize the acid pH within the phagolysosome necessary for activating lysosomal hydrolytic enzymes. Significant changes in the pattern of Brucella proteins during intracellular growth have been recorded (Rafie-Kolpin et al., 1996). As expected, a proportion of the proteins expressed are heat-shock proteins, though others are possibly necessary to deal with other harsh conditions (Lin and Ficht, 1995; Teixeira-Gomes et al., 2000). Neutralization of Oxygen and Nitrogen Intermediates Brucella organisms produce phos-

phomonoesterases, high concentrations of periplasmic cytochromes, Sod and catalase, all proteins that may be involved in the protection of the bacteria against free hydrogen peroxide and superoxide radicals generated by phagocytic cells (Kim et al., 2000; Saha et al., 1990; Tatum et al., 1992). In contrast to the enzyme of some intracellular parasites, such as Legionella or Leishmania, Brucella phosphomonoesterase does not block the production of superoxide anion and does not hydrolyze phosphatidylinositol diphosphate or IP3 molecules (Saha et al., 1990). A collection of Brucella mutants not expressing Cu++/Zn++-SodC or catalase proteins has been generated on the expectation that these defects would increase the sensitivity of the mutants to bactericidal action of oxidative intermediates produced during the respiratory burst (Kim et al., 2000; Tatum et al., 1992). It has been demonstrated that these two enzymes concomi-

The Genus Brucella

403

tantly increase after exposure of the bacteria to peroxide or superoxide ions (Kim et al., 2000), suggesting an adaptive response. In spite of their increased sensitivity to hydrogen peroxide (Kim et al., 2000), the catalase-deficient mutants replicate at the same rate as wildtype bacteria in mice, indicating that, at least in this model, catalase per se does not play a significant role in virulence (Grilló , 1997). Similarly, Brucella SodC-deficient mutants exhibit virulence and establish chronic infections in mice (Latimer et al., 1992). Mutants interrupted in the operon coding for cytochrome bd oxidase, which catalyzes an alternate terminal electron transport step in bacterial respiration, are highly attenuated and unable to replicate in cells. Superexpression of Cu++/Zn++ Sod and catalase in these mutants alleviates the loss of cytochrome bd oxidase (Endley et al., 2001), suggesting that these two enzymes play after all a role in virulence under certain conditions. Role of Heat Shock Proteins Significant changes

in the pattern of Brucella proteins during intracellular growth have been recorded. It has been shown that, in addition to variations in the expression level of 73 proteins, repression of 50 in vitro proteins and induction of 24 new proteins occur during growth of B. abortus within macrophages (Rafie-Kolpin et al., 1996). Acid and oxidative conditions, as well as nutritional and heat stresses, induce the synthesis of “new” bacterial proteins. However, the quantity of these “new” molecules produced in vitro is not equivalent to the amount expressed within macrophages (Lin and Ficht, 1995; Teixeira-Gomes et al., 2000). Brucella htrA, which codes for the periplasmic heat shock-induced serine protease HtrA, is likely to participate in the degradation of oxidatively damaged proteins. Resembling Salmonella and Yersinia htrA mutants, Brucella htrA mutants demonstrate higher sensitivity to oxidative substances, reduced survival in neutrophils and defective replication in macrophages (Elzer et al., 1996; Phillips et al., 1995; Phillips et al., 1997). Comparable to other Brucella stress proteins, such as RecA, the mutation of which also reduces multiplication in mice, htrA Brucella mutants establish chronic infections, suggesting residual virulence (Phillips et al., 1995; Tatum et al., 1992; Tatum et al., 1993). In goats, htrA Brucella mutants demonstrate a more attenuated phenotype (Elzer et al., 1996). Regulated expression of dnaK under oxidative stress seems to be used by bacterial pathogens to withstand the respiratory burst of phagocytes (Caron et al., 1994a). Analysis of Brucella mutants carrying inactivated dnaK and dnaJ (which code for the stress molecular chaperones DnaK and DnaJ) has led to the conclu-

404

E. Moreno and I. Moriyó n

sion that DnaK, but not DnaJ, is required for growth at 37°C. The dnaK mutant survives but does not multiply within phagocytes at a temperature of 30°C, whereas the dnaJ mutant multiplies normally (Köhler et al., 1996). It has been proposed that intracellular Brucella organisms enter into a period of starvation that would favor their resistance to oxidative conditions found within vacuoles (Alcantara et al., 2000; Robertson and Roop, 1999; Robertson et al., 2000a). Brucella abortus hfq mutants (defective in the RNA chaperone host factor (HF-1)) do not replicate in macrophages, but initially they multiply in mice, suggesting that HF-1, normally required for maintenance at stationary phase, is also necessary during intracellular survival. The HF-1 protein participates, within several pathways, in the regulation of the sigma factor RpoS, required for maintenance at the stationary phase (Robertson and Roop, 1999). As a consequence, the deficient hfq mutant displays an impaired stress response and problems for adapting to the stationary growth phase. This mutant, impaired for long-term survival under nutrient deprivation, also demonstrates growth stage- and medium-dependent sensitivity to hydrogen peroxide and a decreased capacity to resist acidic environments, conditions likely to be found intracellularly. Brucella lon mutants are impaired in their capacity to resist hydrogen peroxide and puromycin and display reduced survival in macrophages and significant attenuation in mice during the initial periods, but not at later times. The ATPase-dependent Lon protease is one of the principal enzymes involved in the turnover of stress-damaged proteins. The level of transcription of this protein increases in response to several environmental stresses. It has been proposed that Lon functions as a stress-response protease and is required in Brucella during the initial stages of infection, but it is not essential for the establishment and maintenance of chronic infections in the host (Robertson et al., 2000b). The role of other potential Brucella virulence genes, such as those coding for the heat shock protein GroEL and ClpATPase (Lin et al., 1996), has not been investigated in detail. Brucella suis null mutants for ClpATPase chaperonine behave similarly to the wild-type strain, indicating that ClpA by itself is dispensable for intracellular growth (Ekaza et al., 2000). Role of the Urease Considering ammonium is the principal product and has a strong basic character, the decomposition of urea catalyzed by urease may serve to neutralize the acid pH within the phagolysosome. Urea is in all likelihood a product of the metabolism of Brucella (Response to Environmental Stress) and all Bru-

CHAPTER 3.1.16

cella species but B. ovis have a cytoplasmic urease. However, the urease-defective mutants do not display attenuation and behave similarly to the parental pathogenic strains in some virulence assays, evoking what has been observed with the catalase and SodC mutants (Jubier-Maurin et al., 2001; Grilló , 1997). This does not rule out a possible role for urease during the initial steps of host invasion in the penetration through the intestinal route (Brucella Invades Healthy Hosts). Inhibition of Lysosome-Phagosome Function After 24 h, epithelial cells infected with virulent B. abortus display a significant increase in the number of intracellular bacteria distributed in the perinuclear region corresponding to the endoplasmic reticulum. In contrast, cells treated with killed Brucella harbor a small number of intact bacteria together with bacterial degradation products scattered throughout the cytoplasm, an action that proceeds with time. At these stages, cathepsin D, a well-known marker for lysosomes, colocalizes with vacuoles containing killed B. abortus and bacterial degradation products, attesting that phagosomes have fused with lysosomes (Pizarro-Cerdá et al., 1998a; Pizarro-Cerdá et al., 1998b). By contrast, intracellular virulent Brucella organisms seldom colocalize with cathepsin D in epithelial cells, indicating that this bacterium avoids fusion with lysosomes. In vitro fusion experiments between vacuoles containing Brucella and lysosomes isolated from macrophages show that although compartments containing killed bacteria fuse with lysosomes, vacuoles containing live Brucella do not fuse with these organelles (Naroeni et al., 2001). Moreover, intracellular Brucella organisms do not hamper the fusion of latex beads with lysosomes, a phenomenon that agrees with the cellular parasitism displayed by some rough attenuated Brucella mutants. Some rough intracellular brucellae follow the constitutive endosome-lysosome route, whereas others traffic to the endoplasmic reticulum in the same cell. The key virulence factor is not the general inhibition of lysosome-phagosome fusion in cells, but rather the active modification of the Brucellacontaining compartment, making this organelle non-fusiogenic with lysosomes. It has been found that phagosomes containing live Mycobacterium retain a host protein (TACO, a phagosomal coat protein that prevents degradation of mycobacteria in lysosomes) that prevents delivery of lysosomes into the parasitecontaining phagosome (Ferrari et al., 1999). Similarly, it has been proposed that the Leishmania parasite inhibits phagolysosome biogenesis through insertion of a lipophosphoglycan into the phagosome membrane, precluding in this

CHAPTER 3.1.16

manner the fusion of the phagosome with lysosomes (Desjardins and Descoteaux, 1997). Indirect evidence for inhibition of neutrophil “degranulation” by B. abortus has been obtained from studies of the effects of live or heat-killed organisms and bacterial extracts. Brucella abortus extracts apparently devoid of enzymes and LPS (but containing 5′-guanosine monophosphate and adenine) inhibited neutrophil degranulation (Canning et al., 1986). The same molecules seem to inhibit the myeloperoxidasehydrogen peroxide-halide activity by specifically hampering degranulation of peroxidase-positive polymorphonuclear granules (Bertram et al., 1986). Similarly, studies conducted with B. abortus extracts were capable of inhibiting phagosome-lysosome fusion in macrophages (Frenchick et al., 1985). It has been observed that smooth Brucella LPS enhanced the intracellular survival of rough mutant bacteria in bovine neutrophils (Soto et al., 1991). Despite this, some authors have proposed that LPS does not participate in the inhibition of phagosome-lysosome fusion (Kreutzer and Robertson, 1979a; Frenchick et al., 1985). All these investigations, although valuable, need to explain the fact that only live virulent Brucella is capable of hampering lysosomal fusion (Pizarro-Cerdá et al., 1998c; Pizarro-Cerdá et al., 1998b; Naroeni et al., 2001). Lipopolysaccharide Fails to Stimulate Lysosomal Activity Several studies have cited the Brucella LPS molecule as a virulence factor (Freer et al., 1996; Rasool et al., 1992; Velasco et al., 2000). However, in contrast to what has been proposed for other LPS, the virulence of Brucella LPS is not expressed in the classical form of an endotoxic active molecule (Fig. 18). It is known that LPS from most Gram-negative bacteria activates phagocytic cells by stimulating the respiratory burst, inducing the production of bactericidal nitrogen intermediates and the generation of active cytokines (Brade et al., 1988; Cline et al., 1968; Kelly et al., 1991). All these mechanisms contribute to the destruction of intracellular bacteria and promote the fusion of the ingested bacteria with lysosomes (Silverstein and Steinberg, 1990). In contrast, Brucella LPS practically does not stimulate the oxidative burst or stimulate the release of lysozyme in phagocytic cells and fails to generate significant amounts of bactericidal reactive oxygen intermediates (Jiang and Baldwin, 1993a; Jiang et al., 1993c; Rasool et al., 1992). Furthermore, Brucella LPS induces very little release of interferon gamma (IFNγ) or tumor necrosis factor (TNF), substances which are known to enhance lysosomal fusion and the microbicidal mechanisms of phagocytic cells (Goldstein et al., 1992; Keleti et al., 1974). In this respect, the low biological activity of Brucella

The Genus Brucella

405

LPS may be envisioned as an advantage of Brucella parasites to adapt to intracellular life. Within this context, the Brucella LPS is considered a virulent factor. Iron Chelation and Bactericidal Action B. abortus organisms secrete 2,3 dihydroxybenzoic acid and the catecholic siderophore brucebactin (Denoel et al., 1997a; González-Carrero et al., 2002; Leonard et al., 1997; Ló pez-Goñ i et al., 1992; Uptake of Inorganic Nutrients). Hypothetically, iron-capturing molecules may compete for intracellular iron within the macrophage and inhibit iron-mediated bactericidal killing systems (Leonard et al., 1997). It has been demonstrated that iron-loaded macrophages have enhanced capability to kill or prevent the replication of intracellular B. abortus (Jiang and Baldwin, 1993b). This effect was demonstrated with opsonized and nonopsonized bacteria, as well as with attenuated and virulent strains of B. abortus. The augmented bactericidal activity in the presence of iron may be indirectly mediated by the Haber-Weiss-Fenton and related reactions (Jiang and Baldwin, 1993b). In these reactions, hydroxyl radicals are generated from hydrogen peroxide in the presence of Fe++, which is oxidized to Fe+++. Generation of more hydroxyl radicals and Fe+++ results from the reaction between superoxide and Fe++. The fact that the bactericidal action could be blocked with hydroxyl scavengers supports this general idea. It is likely that, in activated macrophages, the concentration of superoxide ions and hydrogen peroxide increases within the lysosomes, thus providing the substrates for the above-summarized reactions. Similar phenomena have been observed with other Gramnegative and Gram-positive bacteria. Although 2,3 dihydroxybenzoic acid has a comparatively moderate ability to chelate iron, it is able to block the Haber-Weiss-Fenton reaction. This compound protects killing of Brucella mediated by activated macrophages during the first 12 h and increases the number of intracellular brucellae recovered after 48 h of infection (Leonard et al., 1997). Mutations in aroC (the gene coding for chorismate synthase) block chorismate synthesis and therefore these mutants do not produce para-amino benzoic acid, a precursor of 2,3 dihydroxybenzoic acid. Brucella aroC mutants show reduced virulence, supporting a role for this molecule and/or the siderophore during intracellular survival (Foulongne et al., 2001; Hong et al., 2000). However, aroC mutants are pleiotropic, and ∆entC B. abortus mutants (which are defective in isochorismate synthase and thus do not produce 2,3 dihydroxybenzoic acid) are not attenuated in mice (Bellaire et al., 1999). Concurrent with this, B. abortus mutants in an entF homologue do not produce brucebac-

406

E. Moreno and I. Moriyó n

tin or excrete 2,3 dihydroxybenzoic acid but replicate normally in murine macrophages (González-Carrero et al., 2002). Therefore, neither brucebactin nor 2,3 dihydroxybenzoic acid seems to work as a virulence factor in the mouse model. However, there are reports on the attenuation of B. abortus ∆entC in pregnant goats, suggesting that the situation may be different in natural hosts (Bellaire et al., 1998). This is, however, somewhat contradictory to the fact that Brucella bacterioferritin deletion mutants replicate normally within human phagocytes (Denoel et al., 1997a). Modulation of Intracellular Trafficking After phagocytosis, bacteria are found inside a membrane-bound compartment, which undergoes a maturation process that depends upon the nature of the ingested organism (Méresse et al., 1999). The “constitutive route” followed by inert particles, such as latex beads or killed organisms, is revealed by the sequential acquisition of molecules displayed on the vacuole-containing bacteria (Garin et al., 2001). Immediately after uptake, the phagocytic vacuole exhibits plasma membrane molecules, then markers of early endosomes followed by molecules distinctive of late endosomes, and finally elements characteristic of lysosome molecules. Among these molecules, small GTPases of the Rab family, involved in the regulation of fusion events along the endocytic pathway, reflect the capacity of the phagosomes to fuse with endocytic organelles. It has been proposed that phagosome maturation occurs by multiple transient fusion events, a process referred to as “kiss-and-run.” Intracellular pathogens have evolved strategies to avoid this progressive transformation of their vacuole to a phagolysosome, thereby evading killing mechanisms. Brucella organisms, as do other intracellular pathogens, actively evade the constitutive route, promoting their migration to a compartment where they are capable of extracting nutrients, and replicate (Pizarro-Cerdá et al., 1998b). This process is commanded by the interaction of complex mechanisms with the intervention of several genes and molecules. The absence of one or more of these elements alters the intracellular biogenesis of the bacteria and breaks the natural equilibrium established between the parasite and its host cell. Escaping from the Endocytic Pathway Brucella bvrS-

bvrR mutants forced to penetrate CNF-treated HeLa cells follow the degradative pathway (C. Guzmán-Verri et al., unpublished observations), indicating that this two-component, sensory regulatory system is involved not only in penetra-

CHAPTER 3.1.16

tion (Mechanisms of Entry to Host Cells) but also in controlling vacuole maturation. Cyclic glucans of plant pathogenic bacteria are essential factors for parasitism (Breedveld and Miller, 1994). Brucella possess a periplasmic cyclic-β(1,2)-glucan, which is not osmotically regulated, in contrast to what happens in Agrobacterium or Rhizobium parasites (Briones et al., 1997). Brucella cgs mutants, unable to produce cyclic glucan, are avirulent in mice and incapable of avoiding fusion with lysosomes (Iñ on de Iannino et al., 1998; J. P. Gorvel, personal communication). Since Brucella cgs mutants adhere and penetrate cells similarly to the wildtype bacteria, it is concluded that cyclic glucans are necessary for the correct biogenesis of these organisms. Recently, it has been described that a genetic system (virB) encoding for a type IV secretion machinery (Boschiroli et al., 2002; Comerci et al., 2001; Delrue et al., 2001) is also involved in regulating the intracellular trafficking of Brucella. Type IV secretion systems are complex structures composed of several proteins, some of which concomitantly span the inner and the outer-membrane (Fig. 5). These systems are specialized in transferring molecules (proteins or DNA) from the internal to the external milieu of the bacterial cell (Christie and Covacci, 2000). The Brucella type IV secretion system is composed of 13 open reading frames and shows similarity with the VirB complex of other cellassociated bacteria, such as Agrobacterium, Legionella and Rickettsia. Polar mutations in the virB1 and virB10 abolish the ability of Brucella to replicate in mice and cells (Comerci et al., 2001). Nonpolar mutation in virB8 and virB10, coding for internal membrane proteins, generates attenuated bacteria. Mutations in virB12 and virB13 do not demonstrate defects (O’Callaghan et al., 1999; D. J. Comerci, personal communication). Mutations in virB genes do not have an effect in the attachment and internalization of Brucella to cells (Boschiroli et al., 2002; Comerci et al., 2001; Delrue et al., 2001), although some authors claim that the VirB system is implicated in internalization (Watarai et al., 2002). Brucella organisms harboring either polar or nonpolar mutations in virB10 are capable of penetrating cells, as is the wild-type Brucella, localizing in LAMP-1-positive compartments at early times of infection. However, after this period, virB10 polar mutants are sorted to degradative compartments positive for lysosomal markers, whereas nonpolar virB10 mutants remain within compartments devoid of lysosomal or endoplasmic reticulum proteins, but retain early endosome markers. After 12 h, a large proportion of intracellular nonpolar virB10 mutant bacteria are recycled to the cell surface. Once outside, the bacteria seem to replicate adhered to the cell

CHAPTER 3.1.16

envelope. These results indicate that although the virB genes are not required for attachment or invasion, they are necessary for regulating the intracellular trafficking from early endosomes to the endoplasmic reticulum. In addition, the VirB10 product seems to be essential for preventing the fusion of Brucella-containing vacuoles with lysosomes. Therefore, the absence of VirB11 would preclude the correct assembling of the type IV secretion machinery necessary for secreting bacterial substances that prevent lysosomal fusion. It follows that the absence of VirB10 will allow a defective but partially functional secretory system competent for controlling lysosomal fusion, but incapable of releasing substances necessary for promoting retrograde Brucella trafficking to the endoplasmic reticulum. The putative virB10 and B11 genes seem to code for an internal transmembrane protein of unknown function (structural?) and for a cytoplasmic or inner membrane protein that has a conserved Walker A NTP-binding motif, respectively. Gene reporter analysis has revealed that the expression of the Brucella VirB system is activated during the internalization (from 3–12). Then the expression of the system diminishes, corresponding to the replicating time in the endoplasmic reticulum (Boschiroli et al., 2002; Comerci, personal communication). Some authors claim that virB induction only occurs once the bacteria are inside the cells and phagosome acidification has occurred, being this the major signal for expression of this system (Boschiroli et al., 2002). Others, in contrast, sustain that virB operon is turned on during stationary phase and that acidic conditions do not affect the expression of the VirB system (Sieira et al., 2000; Comerci, personal communication). Therefore, it seems that the type IV secretion machinery is required for controlling the brief trafficking of Brucella through the endocytic network until the bacterium reaches its replicating niche within the endoplasmic reticulum. Once the bacterium has reached the endoplasmic reticulum, the secretion apparatus may be turned off. Trafficking from the endocytic network to the endoplasmic reticulum via autophagosomes requires not only live virulent bacteria, but a physiologically balanced Brucella-containing vacuole. For instance, in cells expressing an activated form of rab5 (bound to GTP), an important fraction of the internalized parasites are unable to escape from the early Brucellacontaining compartment (Chaves-Olarte, 2002). Although a fraction of bacteria are capable of reaching the endoplasmic reticulum in these mutant cells, unexpectedly a few Brucella escape from these “early” giant vesicles to the cytoplasm (J.-P. Gorvel, personal communication). These events suggest that Brucella is capable of

The Genus Brucella

407

adapting to different intracellular environments by controlling its own intracellular trafficking through direct interaction with its host vacuole. Escaping from Autophagosomes The first step in the biogenesis of autophagosomes is the acquisition of lysosomal membrane-associated proteins. Acidification of the maturating compartment occurs by inclusion of the H-ATPase, and finally delivery of acid hydrolases allows the degradation of intravacuolar isolated cytoplasmic materials (Dunn, 1994). In fact, it has been shown that nocodazole treatment causes the accumulation of acidic autophagosomes that lack acid hydrolases, supporting the concept that vacuole acidification and acquisition of hydrolytic enzymes are separate events. The presence of LAMP-1 and LAMP-2 but the absence of cathepsin D at 2 h postinoculation in B. abortus-containing phagosomes also supports the model of a stepwise maturation of autophagic vacuoles. Whether B. abortus-containing autophagosomes are able to acquire the H-ATPase remains to be established. Brucella phagosomes seem to acidify rapidly after infection in murine J774 macrophages (Arenas et al., 2000; Porte et al., 1999). This acidification step seems to be independent of phagosome-lysosome fusion. In Vero cells, repressors of endosome acidification, such as chloroquine and ammonium chloride, reduce the number of Brucella with respect to controls (Detilleux et al., 1991). Adjustment of intravacuolar pH has been shown as essential for the activation of virulence genes in certain intracellular parasites. Parasitophorous vacuoles of Leishmania amazonensis maintain an acidic pH in infected macrophages (Antoine et al., 1990). New sets of proteins are synthesized by S. typhimurium upon infection of host cells after modulation of the phagosome pH (Buchmeier and Heffron, 1990). This latter bacteria induces reduction of phagosome acidification to activate the virulence genes of the phoPphoQ complex (Alpuche-Aranda et al., 1992). Brucella abortus is able to synthesize a new set of proteins during macrophage infection (Lin and Ficht, 1995; Rafie-Kolpin et al., 1996), suggesting that intracellular conditions (one of which could be intravacuolar pH) may contribute to the activation of genes necessary for trafficking from autophagosomes to the endoplasmic reticulum (Pizarro-Cerdá et al., 1998c). The critical difference between the attenuated vaccine strain 19 and the virulent B. abortus 2308 seems to be the inhibition of autophagosome maturation by the virulent strain (Fig. 36). The attenuation of strain 19 may lay in its incapacity to respond to environmental stimuli present in the autophagosome (acidification, for example) that could activate virulence

408

E. Moreno and I. Moriyó n

genes for the expression of proteins important to remodel the autophagosome. Life Within the Replicating Niche In addition to a

suitable source of carbon and energy, Brucella requires some cofactors and vitamins for growth (Nutritional Requirements). It is also able to use some amino acids as a source of carbon and energy and to eliminate the urea that could build up as a metabolite noxious for the host and the bacterium under some conditions (ATP Synthesis and Respiratory Chain; Oxidation of Amino Acids). Besides being a strategy for avoiding lysosome fusion during later steps of intracellular invasion, association of pathogenic Brucella with the endoplasmic reticulum could be a strategy for obtaining metabolites synthesized or translocated to this compartment (Simon and Blobel, 1991). By stimulating host cell autophagy, the bacteria would increase degradation of host proteins and availability of amino acids. However, the mechanism by which these small compounds would be delivered back to the endoplasmic reticulum and used by the bacteria should be defined. In contrast to professional phagocytes, it seems that the placental environment offers a privileged site for brucellae replication. It is becoming evident that trophoblasts are metabolically active cells capable of producing a variety of hormones and other factors which may stimulate the growth of Brucella organisms (Enright and Samartino, 1994). Genes coding for putative efflux pumps capable to serve as export of quorum sensing metabolites have been identified in Brucella (DelVecchio et al., 2002). Quorum sensing hydrophobic molecules such as acyl-homoserine lactone released by Brucella down regulate the expression of virB genes in vitro (Taminiau et al., 2002), suggesting that this small hydrophobic molecule could play a role in the control of the type IV secretion system, necessary for the intracellular trafficking, but unnecessary and inhibitory during intracellular growth inside the endoplasmic reticulum. Role of the Outer Membrane in Nutrient Uptake

The overall higher hydrophobicity of Brucella cell envelopes, the close association among the macromolecules of the outer-membrane and the properties of the porins are the factors implicated in the selective penetration of nutrients inside the bacterial cell. It has been demonstrated that the absence of a barrier to hydrophobic substances is linked to the structure of the Brucella LPS core and lipid A (Freer et al., 1996; Martínez de Tejada and Moriyó n, 1993; Velasco et al., 2000). The possible advantages of a hydrophobic envelope for intracellular α-2 subclass

CHAPTER 3.1.16

Proteobacteria also have come to light by the finding that Rhizobium LPS becomes highly hydrophobic during bacteroid development (Kannenberg and Carlson, 2001). The net result of this structural change in the LPS is that intracellular bacteroids have a more hydrophobic outer-membrane than the free-living rhizobiae have. This adaptive condition could promote the exchange of nutrients and favor intracellular life of the bacteroids. Obvious comparisons between the intracellular lifestyle of Brucella and Rhizobium emerge, since these two bacteria are phylogenetically close relatives (Moreno et al., 1990). The permeability of the Brucella outermembrane to sexual hormones and siderophores is discussed elsewhere in the chapter (The Hydrophobic Pathway; Uptake of Inorganic Nutrients). As noted (Porin Proteins), B. abortus only expresses one of the two porin protein genes it carries, and it has been suggested that the second one may be specifically expressed during intracellular parasitism (Marquis and Ficht, 1993; Ficht et al., 1989). Auxotrophic Intracellular

and Cell Cycle Genes During Life Many of the mutations

described in this section may hamper the adequate extraction of nutrients from the replicating niche. It is likely that these genes are devoted to the regulation of vital functions not directly involved in virulence, but with more general aspects of the Brucella physiology. Mutants displaying reduced virulence and/or reduced intracellular survival within macrophages have been identified by signature-tagged transposon mutagenesis (Foulongne et al., 2000). Several of these attenuated mutants carry auxotrophic defects, such as those necessary for leucine, arginine or aromatic acid biosynthesis, whereas others carry deficiencies in the synthesis of chorismate for the generation of para-aminobenzoic acid necessary for quinone synthesis, 2,3 dihydroxybenzoic acid, brucebactin and folic acid. Mutations in genes involved in the glucose metabolism, such as in the gene coding for phosphoglucose isomerase, also attenuate Brucella, probably owing to several pleiotropic defects, including the synthesis of cell wall peptidoglycan. Similarly, transposon insertion in the gpt-like gene coding for hypoxanthineguanine biosynthesis attenuates Brucella, probably owing to alterations in the nucleotide biosynthesis. In support of this are the B. melitensis purE (purine auxotrophic) mutants that display reduced growth in macrophages (Cheville et al., 1996a; Drazek et al., 1995). The purE mutation has minimal effect on internalization, but effectively blocks intracellular replication. The purE mutation may cause bacterial death simply as a result of starvation.

CHAPTER 3.1.16

Other mutants, also identified by signaturetagged mutagenesis (Foulongne et al., 2000), have defects in regulatory systems, such as LysR transcriptional regulator, which is part of a positive regulator system of virulence genes in several bacteria. Mutants in the expression of NtrY protein (a sensor of an Ntr-related regulator; Dorrell et al., 1999) are weakly attenuated, probably as a result of a pleiotropic negative effect on the ntr regulon (The NtrBC System). As in other bacteria, mutations in genes involved in glutamine metabolism reduce the ability of Brucella to replicate in macrophages (Foulongne et al., 2000). The Brucella ccrM gene codifies for a CcrM DNA methyltransferase that catalyzes the methylation of the adenine in sequences GANTC (Methylation and Control of Chromosome Duplication and Cell Cycle). This gene performs important functions during cell division and it is, therefore, essential for viability (Robertson et al., 2000b; Wright et al., 1997). Increase in the ccrM copy number, in addition to altering the morphology of the bacterial cells, attenuates Brucella, indicating that controlled cell cycle and bacterial division are necessary for intracellular survival. Mutations in the bacA gene, which codes for a putative cytoplasmic membrane transporter, render B. abortus avirulent (LeVier et al., 2000). Brucella bacA mutants display altered transport of molecules and then deficient Brucella replication within cells. Auxotrophic mutants for ferrochelatase, the enzyme involved in the last step of the heme synthesis (catalyzing the incorporation of ferrous iron into the protoporphyrin molecule), are also attenuated (Almiró n et al., 2001). Maintaining the Host Cell Alive Some parasites may promote programmed cell death, whereas others are prone to prevent it, prolong cell life, or are even capable of stimulating replication of their host cells. Proteobacteria of the α subdivision, such as Rickettsia, Agrobacterium, Rhizobium and Brucella, as well as other intracellular parasites capable of establishing chronic infections (i.e., Mycobacterium and Chlamydia) prevent cell death. Heavily infected Brucella trophoblasts, epithelial cells or macrophages do not display signs of necrosis or apoptosis (Anderson et al., 1986b; Anderson et al., 1986c; Detilleux et al., 1990b; Jiang and Baldwin, 1993a; Tobias et al., 1993; Fig. 30). DNA synthesis, microtubule spin formation, chromosome migration, karyokinesis and cytokinesis are not inhibited by intracellular Brucella (Chaves-Olarte et al., 2002). As a consequence, dividing cells filled with brucellae are frequently observed in vivo and in vitro (Detielleux et al., 1990a; Detilleux et al., 1990b; Detilleux et al., 1991). In CNF-treated HeLa

The Genus Brucella

409

cells, cytokinesis is inhibited due to paralysis of actin filaments without affecting nuclear division. When these cells are infected with Brucella, karyokinesis proceeds without signs of degeneration, despite the large number of intracellular Brucella within the endoplasmic reticulum (Chaves-Olarte et al., 2002). Live but not killed B. suis prevent programmed cell death of infected human monocytes (Gross et al., 2000). This suggests that infection protects host cells from several cytotoxic activities generated during the immune response. Since both invaded and noninvaded cells are guarded against apoptosis, it has been suggested that this protective mechanism is mediated through soluble substances released during bacterial infection. Apoptosis inhibition is independent of LPS and requires the overexpression of the A1 gene by infected cells, a member of the bcl-2 family involved in the survival of blood-forming cells. In contrast to other LPS, Brucella LPS practically does not display endotoxicity and is a poor inducer of cytotoxic mediators (Goldstein et al., 1992; Ló pez-Urrutia et al., 2000; Moreno et al., 1981; Rasool et al., 1992). This property shared by other intracellular animal pathogens (such as Rickettsia, Legionella and Bartonella) may be envisioned as an evolutive advantage of Brucella parasites for adaptation to intracellular life (Freer et al., 1996; Moreno, 1992; Rasool et al., 1992; Velasco et al., 2000). It is not surprising that lipid As of L. pneumophila and B. abortus parasites have coevolved to have a similar structure and, in consequence, display low endotoxicity and reduced ability to stimulate cells (Goldstein et al., 1992; Ló pez-Urrutia et al., 2000; Moreno et al., 1981; Rasool et al., 1992; Zähringer et al., 1995; Fig. 7). This property may be idiosyncratic and useful for intracellular bacteria. Modulation of the Immune Response Once inside macrophages, pathogens might diminish or abrogate their antigen presentation capacity, thus reducing the T cell-mediated immune responses. The LPSs from different bacteria have been shown to modulate the immune responses in several systems (Cella et al., 1997; Knolle et al., 1999; Krieger et al., 1985; Uchiyama et al., 1984). Brucella abortus LPS molecules accumulate inside lysosomal compartments and associate with MHC-II proteins in antigen-presenting cells (Forestier et al., 1999a; Forestier et al., 1999b; Forestier et al., 2000). The intracellular LPS, which remains for long periods without being degraded, is exported to the cell surface where it forms stable macrodomains (Figs. 35 and 37). Once inside macrophages, B. abortus LPS impairs the MHC-II presentation pathway, but not MHC-I presentation of foreign peptide antigens (Fig. 37). This impairment is neither

410

E. Moreno and I. Moriyó n

CHAPTER 3.1.16

A

HEL or Peptide 34-45 B

% of cell activation

100

75 HEL Peptide 34-45

50

25

0 - LPS

+ LPS

Fig. 37. Brucella LPS binds to MHC-II molecules and inhibits T cell responses to protein antigens. A) Brucella LPS (red) and MHC-II (green) colocalize (yellow) within compartments and on the surface of murine macrophages. B) Inhibition of antigen presentation of hen’s egg lysozyme (HEL) peptides and derived synthetic peptide (34–45) to respective specific T CD4+ clones by Brucella LPS-treated macrophages. Colocalization was detected by double immunofluorescence and viewed in a confocal microscope. Notice that Brucella LPS-MHC-II complexes establish large macrodomains (white arrow points to area of yellow fluorescence) on the surface of phagocytic cells. Adapted from Forestier et al. (2000). Courtesy of Dr. J-P Gorvel, CIML, MarseilleLunimy, France.

from a deficient uptake or catabolism of the native antigen nor from a reduced MHC-II surface expression, reduced number of B7 membranous costimulatory molecules, or defective α/ β dimer formation. In addition, this inhibitory effect is not due to a direct suppressive action of LPS on T cells, independent of macrophages

(Forestier et al., 2000). Brucella LPS macrodomains at the macrophage plasma membrane are highly enriched in MHC-II molecules (Fig. 37), suggesting that the LPS-MHC-II macrodomains may impair the appropriate recognition of protein peptide-MHC-II complexes by CD4+T cells (Forestier et al., 2000). The presence of such MHC-II-LPS macrodomains does not prevent the binding of antigen peptides into the groove of MHC-II molecules. Therefore, the Brucella LPS-induced interference on MHC-II antigen presentation is likely to occur distal to intracellular events leading to the meeting of antigenic peptides and MHC-II molecules. The LPS molecules embedded in the membrane of MHC-IIpositive compartments may interact with already peptide-MHC-II forming ternary complexes, which then recycle to the plasma membrane. In one model, the LPS O-polysaccharide, facing the external milieu, could prevent the accessibility of MHC-II complexes to their specific T cell receptor. Another model is related to a superantigenlike function in T cell activation. It is known that superantigens modify the geometry of TCRpeptide/MHC-II complexes, which may be less critical for T cell activation than certain other factors, in particular those involved in the stability of the resulting complex (Andersen et al., 1999; Kersh et al., 1998). The serial triggering (Itoh et al., 1999; Valitutti et al., 1995; Viola and Lanzavecchia, 1996) and kinetics proofreading models (Rabinowitz et al., 1996) of T cell activation suggest that the short half-lives of TCRpeptide/MHC-II complexes are required for efficient T cell stimulation. The presence of Brucella LPS-MHC-II macrodomains in macrophages has been detected even after 60 days, thus highlighting the remarkable stability of these surface LPS macrodomains. In this model, LPS could downregulate T cell responses by stabilizing the MHC-II/peptide complexes at the cell surface. Consequently, in contrast to super antigens, Brucella LPS would be less efficient at triggering T cells because they form TCR-LPSMHC-II complexes with a very long half-life. The in vitro inhibition of the immune response correlates to that observed in vivo upon infection by Brucella. It is worth noting that chronic brucellosis is accompanied by a general immunosuppression that can be revealed by using an IL-2 detection system (Zhang, 1992; Zhang et al., 1993; Forestier et al., 1999a). Infected macrophages may exert a negative feedback, diminishing lymphocyte proliferation in response to Brucella antigens (Cheers et al., 1980; Riglar and Cheers, 1980). It has been proposed that chronically infected macrophages may fail to act as targets of T cells and may downregulate T lymphocyte function (Baldwin and Winter, 1994). Moreover, it is known that Brucella inhibits the production

CHAPTER 3.1.16

of TNFα of infected phagocytes by releasing a protease-sensitive inhibitor for the expression of this cytokine (Caron et al., 1996). Therefore, it is reasonable to speculate that intracellular Brucella uses this strategy to regulate the activation of phagocytic cells and promote in this manner its own survival within these cells. Since LPS is also released from bacteria inside host cells (Fig. 35) and this molecule is not degraded by peritoneal macrophages, it can be hypothesized that one additional form by which Brucella may contribute to immunosuppression is the release of LPS inside cells (Forestier et al., 1999b). In this respect, Brucella LPS, together with other molecules, may play a central role in the immunosuppression observed upon brucellosis infection and may account for the presence of anergic T cells in infected patients (Renoux and Renoux, 1977). Experiments have demonstrated that Brucella is capable of infecting small lymphocytes in the cortical zone of lymph nodes (Cheville et al., 1996a). The infected lymphocytes harboring intracellular bacteria within vacuolar compartments with no signs of degradation look normal (Fig. 30). At the present time, the implications of lymphocyte Brucella parasitism are not known. The fact that Brucella LPS complexes with MHC-II in these cells (Forestier et al., 1999a) opens the possibility that LPS released by intracellular Brucella could modulate antigen presentation in lymphocytes, as it does in macrophages (Forestier et al., 2000).

Control of the Infection Strong humoral and cellular immune responses are generated during Brucella infection. Although the role of cell mediated immune response has been regarded as the essential mechanism for controlling the infection, antibodies may play a more or less relevant role, depending upon the specie of the infected animal. Comprehensive reviews can be found in references (Baldwin and Winter, 1994; Liautard et al., 1996; Nicoletti and Winter, 1990b; Oliveira, et al., 1998). Antibody Response Brucella infections generate an elevated production of antibodies. The most relevant molecules involved in the humoral response of animals or humans infected or vaccinated with smooth strains are by far the LPS and the NH polysaccharides (Jones et al., 1980; Marín et al., 1999; Moreno et al., 1984a; Nielsen et al., 1988; Spink, 1956). Mainly at early stages of the infection (first month), it has been estimated that close to 80% of the antibodies against Brucella are directed toward the O chain and NH determinants, with the great majority of the immunoglobulins being against the C epitopes

The Genus Brucella

411

(in smooth strains) (Fig. 34). Moreover, most of the IgG1 passively transferred through colostrum to newborn calves are directed against these two bacterial surface molecules (Antibody Detecting Tests in Infected Patients; Smoothlipopolysaccharide Tests in Infected Animals). Antibodies recognizing core and lipid A determinants (Fig. 34) are not detected by conventional serological methods in the sera from infected or vaccinated bovines with smooth strains, unless purified antigens are used. In contrast, individuals infected or vaccinated with rough brucellae generate a strong antibody response against LPS core and lipid A determinants as well as toward Omps (Bowden et al., 1995; Gamazo et al., 1989; Riezu-Boj et al., 1990; Rojas et al., 1994). The most immunogenic proteins are the Omps of group 3 (Omp3a and Omp3b and Omp31), some of which are strongly bound to the LPS (Gamazo and Moriyó n, 1987; Kurtz and Berman, 1986; Riezu-Boj et al., 1990; Rojas et al., 1994), and stress response proteins (GroEL, DnaK, Sod, PurE and HthA) (Protein Tests in Infected Patients; Protein Tests in Infected Animals). Antibodies against various cytoplasmic soluble proteins, ribosomal proteins, periplasmic proteins, outer-membrane lipoproteins and porins, as well as against peptidoglycan, have also been detected (Fig. 38). Most of the antibodies against proteins appear at later stages (after the third week) of the infection (Protein Tests in Infected Patients). The immunoglobulin profiles of the anti-LPS response in newly infected bovines, humans, goats and mice measured by ELISA or discriminatory agglutination are characteristic of a Tdependent cell response (Fig. 39), despite the fact that Brucella LPS, as well as killed organisms, has been defined as a T-independent antigen (Kurtz and Berman, 1986; Moreno and Berman, 1979; Moreno et al., 1984b; Rennick et al., 1983; Tenay and Strober, 1985). The secondary antibody responses observed in antibiotictreated individuals who have relapsed are also typical of a T-dependent response with strong generation of IgG1 antibodies (Antibody Detecting Tests in Infected Patients; Smoothlipopolysaccharide Tests in Infected Animals). Under natural conditions, IgG1 antibodies may remain for life (Castañ eda, 1961; Foz et al., 1954; Spink, 1956). This may be due to the persistence of the bacteria in tissues (Enright, 1990b; Enright, 1990a; Spink, 1956) or alternatively to the permanence of Brucella LPS within phagocytic cells for long periods. In support of this last statement are the findings of LPS within macrophages of infected mice after several months (Modulation of the Immune Response) and the detection of large quantities of apparently intact LPS in granulomatous lesions of individuals with

412

E. Moreno and I. Moriyó n

H

S1

S2

Fig. 38. Crossover immunoelectrophoresis with protein antigens for the diagnosis of brucellosis. The wells on the left contained the sera from a healthy donor (H) and from two patients after a brief (S1) or a prolonged (S2) period of infection before treatment. For reference of the antibody response to LPS see Fig. 39. Courtesy of R. Díaz, University of Navarra, Spain.

chronic brucellosis (R. Díaz, University of Navarra, Spain, personal communication). The antibody response of mice immunized with purified LPS generates a non-typical Tindependent response with relatively large quantities of IgG3 against the O chain and moderate quantities of IgG1 and IgM (Kurtz and Berman, 1986; Moreno et al., 1984b). In contrast to other types of LPS, Brucella LPS is immunogenic for the endotoxin-resistant C3H/HeJ mice and displays a strong IgG1 adjuvant effect in this and other strains (Kurtz and Berman, 1986; Moreno and Berman, 1979; Moreno et al., 1984b). The role of antibodies against Brucella infection has been a matter of controversy for many years. In mice, it is clear that antibodies play an important part in anti-Brucella immunity (Araya et al., 1989), mainly because of the exacerbation of the phagocytosis and bactericidal action mediated by professional phagocytes (Opsonization and Complement Susceptibility). Nevertheless, in other species of mammals, including humans, infections seem to course unaffected in the pres-

CHAPTER 3.1.16

ence of large quantities of circulating antibodies (Ariza, 1998; Enright, 1990a; Enright, 1990b; Shuterland and Searson, 1990). However, maternal antibodies seem to play a role in protecting calves and piglets born within herds with prevalent brucellosis as demonstrated by experimental and field experience (Cunningham, 1977b; Halliday, 1968; Hoerlin, 1957; Plommet, 1977). In addition, whether antibodies to LPS, or to any other antigen, hamper the ability of the bacteria to infect animals under field conditions needs to be examined. The truth is, that despite the current opinion that antibodies do not play a role in human or ungulate infections, at the present this asseveration is not sustained by clear-cut experimental data. This doubt is not trivial and it has practical implications related to the use of vaccines derived from rough strains for controlling brucellosis (Samartino et al., 2000; Schurig et al., 1991). Finally, the role of anti-LPS immunoglobulins either as blockers of antigen recognition or as facilitators of cell invasion also needs to be reexamined in light of the recent advances in our knowledge of Brucella parasitism and intracellular trafficking of bacterial antigens. T Cell Responses In a series of elegant investigations, Mackaness (1967) demonstrated more than three decades ago that acquired cellular resistance could be specifically induced by Brucella organisms or by other facultative intracellular parasites, but nonspecifically expressed for a given period of time. That means that, at some intervals after B. abortus invasion, the Brucellainfected mice will clear Listeria as though they have been immunized against this Gram-positive bacterium. The postulated cellular episodes mediated by lymphocyte cytokines capable of activating macrophages, that eventually could kill both organisms, were the basis for explaining the immunological events necessary to control Brucella inside macrophages (Mackaness, 1967). Passive transfer of CD4+, CD8+ cells and antibodies has clearly demonstrated the role of each of these elements in the control of murine brucellosis (Araya et al., 1989; Araya et al., 1990; Eze et al., 2000). It has been hypothesized that CD8+ cells, in addition to producing IFNγ, could lyse Brucella infected macrophages and downregulate the production of cytokines by Th2 cells, such as IL-10 and the cytokine-like CD8+ T cell antiviral factor (Oliveira et al., 1998). Despite the role played by antibodies and CD8+, it is generally accepted that the ultimate brucellacidal activity in most animals is based on CD4+ type-I immune response, through the generation of cytokines capable of activating macrophages (Baldwin and Winter, 1994; Liautard et al., 1996; Oliveira et al., 1998). In consequence, a more robust respiratory burst, a more powerful pro-

CHAPTER 3.1.16

The Genus Brucella

Primary and secondary antibody response of a cow vaccinated and infected with B. abortus

Antibody response after antibiotic treatment of a human patient followed by a relapse End of treatment Relapse

Antibody titre

IgG 1000 IgA 100 IgM 60

180 Days

300

Optical density 405 nm

S19 vaccinated B. abortus field strain

10000

10

2.25

0.75

0

IgG IgG1 IgG2 IgM

0 75

150 225 Days

300

375

180 Days

60

300

460

Primary and secondary antibody response of S19 infected mouse reinfected with 2308

Optical density 405 nm

Optical density 405 nm

1.6

0

IgG2 IgM IgA

600

Infection

0.8

IgG1

1.50

Primary and secondary antibody reponse of a goat vaccinated and infected with B. melitensis

Rev 1 vaccination

413

B. abortus S19

B. abortus 2308

1.2 0.8 IgG IgG1 IgG2b IgM

0.4 0 0

30

60 Days

90

120

Fig. 39. ELISA immunoglobulin isotype reactivity against B. abortus LPS from humans and animals infected with Brucella. A) Antibody response (over time) of a human patient infected with B. melitensis after antibiotic treatment, followed by relapse. B) Primary and secondary antibody responses of a cow vaccinated with B. abortus S19 and infected with B. abortus 2308, respectively. C) Primary and secondary antibody responses of a goat vaccinated with B. melitensis Rev. 1 and infected with B. abortus, respectively. D) Primary and secondary antibody responses of a mouse infected with B. abortus S19 and reinfected with B. abortus 2308, respectively. Notice the strong IgG (IgG1) response against Brucella LPS, characteristic of a type-2 T-dependent immune response. Since most of antibodies produced are directed against the common Brucella epitopes (Fig. 34), B. abortus LPS antigen bound to the ELISA plates serves its purpose, regardless of whether the individuals were infected or vaccinated with B. abortus or B. melitensis. The profile of the infected patient (panel A) was supplied by Dr. Javier Ariza, from the Infectious Disease Service, Hospital de Bellvitge, Barcelona, Spain.

duction of microbicidal reactive oxygen intermediates, and a more efficient phagosome-lysosome fusion are performed by the activated macrophages (Gross et al., 1998; Jiang and Baldwin, 1993a; Jiang and Baldwin, 1993b; Zhan and Cheers, 1995; Zhan and Cheers, 1996). As expected, the most relevant antigens involved in the T cell responses are bacterial proteins, as revealed by delayed-type hypersensitivity reactions in infected individuals with a mixture of protein extracts (Cell-mediated Immunity Tests). However, nonprotein antigens also may be involved in T cell responses (Bertotto et al., 1993;

Ottones et al., 2000a; Ottones et al., 2000b; J.-P. Gorvel and E. Moreno, unpublished results). Similar to what has been observed with other intracellular bacteria, such as Mycobacterium, Listeria and Francisella, the blood of humans infected with Brucella demonstrates an important increase in γδ T cells (Barnes et al., 1992; Bertotto et al., 1993; Jouen-Beades et al., 1997; Poquet et al., 1998). The T lymphocyte population that specifically expands is one carrying the Vγ9Vδ2 T cell receptor, suggesting that this subset of lymphocytes is generated in response to intracellular pathogens. The γδ T cell repertoire

414

E. Moreno and I. Moriyó n

responds to natural and synthetic low molecular weight nonpeptide ligands of diverse structure, complicating the nature of the recognition pattern carried out by this repertoire. For instance, when stimulated with isopentyl pyrophosphate from M. tuberculosis, Vγ9Vδ2 T cells polyclonally expand and secrete high levels of TNFα and INFγ, without stimulating αβ or Vγ1Vδ2 subsets. Similarly to other intracellular bacteria, Brucella stimulates the production of TNFα and INFγ by Vγ9Vδ2 T cells (Ottones et al., 2000a; Ottones et al., 2000b). It has been shown that Brucella-activated Vγ9Vδ2 lymphocyte subset is capable of impairing the intracellular multiplication. This effect seems to proceed by two different mechanisms. The first one is through the release of a low molecular weight soluble factor that stimulates the production of TNFα and INFγ cytokines, which activate the brucellacidal mechanisms of professional phagocytes. The second mechanism seems to take place by a direct contact-dependent cytotoxic effect on monocytic cells infected with Brucella organisms. The bactericidal activity mediated by this γδ lymphocyte subset during early stages of infections by intracellular bacteria, such as Mycobacterium, Listeria and Brucella, may explain, in part, why acquired cellular resistance could be specifically induced by one intracellular bacteria, but nonspecifically expressed for a given period of time, as indicated at the beginning of this subsection (Mackaness, 1967). Interleukins in Brucellosis Several studies have highlighted the role of IFNγ, TNFα, interleukin (IL)-6, IL-1β, IL-10, IL-12, granulocyte macrophage colony-stimulating factor (GM-CSF), and granulocyte colony-stimulating factor (G-CSF) in host resistance against bacterial infections (Fernández-Lago et al., 1996; Flesh et al., 1994; Flynn et al., 1995; Liautard et al., 1996; Sarmento and Appelberg, 1996; Tanaka et al., 1995). Delayed-type hypersensitivity response of Brucella-infected animals and humans has been used as a diagnostic parameter for brucellosis, as well as an indicator for cell-mediated immune response and release of cytokines in brucellosis (Nicoletti and Winter, 1990b). Particularly, during Brucella-infection IFNγ, TNFα, IL-2, IL-10, and IL-12 seem to control the intracellular growth of Brucella strains within macrophages, whereas IL1α, IL-4, IL-6 and GM-CSF do not have clear effects (Fernández-Lago et al., 1996; Jiang and Baldwin, 1993a; Ottones et al., 2000a; Ottones et al., 2000b; Zhan and Cheers, 1995). Among the various cytokines, IFNγ is the most relevant for generating macrophages with strong brucellacidal activity. Moreover, IL-2, IL-10 and IL-12, cytokines that influence the acquired cellular resistance and specifically contribute to control

CHAPTER 3.1.16

the brucellae multiplication, seem to work via the IFNγ-dependent pathway. It is known that Brucella LPS is a poor inducer of cytokine production, as well as an inefficient activator of phagocytic cells (Goldstein et al., 1992; Ló pezUrrutia et al., 2000; Rasool et al., 1992). Similar to what has been observed with Legionella pneumophila LPS, it seems that the CD14 molecule does not serve as receptor for Brucella LPS (Neumeister et al., 1988; E. Moreno, unpublished observations), precluding the activation of macrophages by this mechanism. This coincidence between two LPS from phylogenetically distant intracellular bacteria does not come as a surprise, since the lipid A structures of these two molecules have co-evolved to the point that they chemically resemble each other (Fig. 7). As an overall conclusion, it seems unlikely that macrophage activation and secretion of IL-12 occur via the Brucella LPS molecule (Huang et al., 1999). The role of TNFα in brucellosis is not completely clear. In contrast to what has been observed in murine macrophages, Brucella strains do not induce TNFα in human macrophages and live Brucella is capable of inhibiting the production of this cytokine (Caron et al., 1994b; Caron et al., 1996). However, pretreatment of human macrophages with exogenous TNFα significantly inhibits the rate of Brucella intracellular replication. TNFα may not be essential for the induction of acquired cellular resistance but it is likely to directly activate effector cells by limiting the multiplication of intracellular Brucella (Ottones et al., 2000a; Ottones et al., 2000b; Zhan and Cheers, 1996). It has been demonstrated that bactericidal action of activated macrophages and expression of macrophage-specific cytokines depend upon the expression of NF-IL6 (Akira et al., 1990; Katz et al., 1993; Lowenstein et al., 1993; Scott et al., 1992; Tanaka et al., 1995). Upon activation of NF-IL6 knockout macrophages by IFNγ, induction of the transcription of TNFα, IL-6, IL1β, GM-CSF, M-CSF, IL-10, and IL-12 is comparable to that observed in normal mice (Tanaka et al., 1995). Strikingly, no induction of G-CSF expression is observed in NF-IL6 knockout mice, being this defect is restricted to macrophages and fibroblasts (Tanaka et al., 1995). Moreover, NF-IL6 knockout mice display a high susceptibility to Salmonella and Listeria infections, suggesting that NF-IL6 plays a role in controlling intracellular parasites (Tanaka et al., 1995). Attenuated B. abortus strain 19 is capable of replicating in NF-IL6 knockout murine macrophages. The levels are comparable to those observed in normal macrophages infected with the virulent Brucella strains (Pizarro-Cerdá et al., 1998c). The role of NF-IL6 in the inhibition of intracellular bacterial replication is related to

CHAPTER 3.1.16

its control of endocytosis and membrane fusion between endosomes and Brucella-containing phagosomes. Addition of G-CSF, where production is impaired in NF-IL6 knockout macrophages, restores both endocytosis and the morphology of endosomes, together with the bactericidal activity. During Brucella infection, it has been observed that endocytosis but not recycling is affected in NF-IL6 knockout macrophages, suggesting that G-CSF promotes fusion events in early phagosomes (Pizarro-Cerdá et al., 1999a; Fig. 35). In NF-IL6 knockout macrophages, NO synthetase is not impaired (Tanaka et al., 1995). The production of reactive oxygen intermediates is, however, lower in NF-IL6 than that in wildtype macrophages (Tanaka et al., 1995), suggesting that NF-IL6 may control the expression of other elements of the respiratory burst. Indeed, it is known that G-CSF enhances the respiratory burst in phagocytes (Yuan et al., 1993). The NO synthetase was found to be associated to intracellular membrane vesicles different from lysosomes and peroxisomes in murine macrophages (Vodovotz et al., 1995). These vesicles could translocate to Brucella-containing phagosomes in normal macrophages and be hampered in NFIL6-deficient macrophages due to the lack of fusion between endosomes and phagosomes. Therefore, one hypothesis is that G-CSF, by completely restoring endosome-phagosome fusion, allows elements of the respiratory burst present in endocytic compartments to reach the Brucella-containing phagosomes and thus to partially re-establish the bactericidal activity of macrophages (Pizarro-Cerdá et al., 1999a). Under these conditions, attenuated Brucella could be targeted to lysosomes and killed, whereas virulent bacteria could still replicate but to a lesser extent than in resting macrophages (Jones and Winter, 1992).

Diagnosis Brucellosis causes a great variety of clinical symptoms in humans, and although a thorough anamnesis (by questioning feeding habits, occupation and recent animal contacts) is very informative, a definite diagnosis cannot be made solely on these bases. A similar situation happens in animal brucellosis: abortion and infertility are the main symptoms, but they are not exclusively caused by Brucella and do not necessarily occur in all infected animals. The laboratory diagnosis of brucellosis logically follows clinical or epidemiological suspicion of the disease and uses direct and indirect tests. Direct tests are those that show the presence of viable bacteria by bacteriological culture or of some molecules specifically produced by the

The Genus Brucella

415

pathogen. They have the advantage that, when positive, the diagnosis is conclusive. However, they often lack sensitivity (i.e., they yield false negative results) and are cumbersome to perform. Indirect tests are those that detect the presumed effects of Brucella infections, which means almost always an analysis of the adaptive immune response. Depending on the tests, they can be very sensitive. However, it has been proven that a proportion of infected animals born to infected mothers do not develop a detectable immune response immediately after birth and that animals that have been infected for a long time may have no significant levels of antibody to the commonly used Brucella antigens. Indirect tests may also lack specificity (i.e., they may produce false positive results). This is so because a positive result of immunological tests can mean not only an active infection, but also the existence of antibodies or memory cells persisting after vaccination or recovery, contact with Brucella not followed by disease, or exposure to immunologically crossreacting bacteria. Thus, without a good knowledge of the circumstances outlined in the previous paragraph, the results of laboratory tests are not always meaningful. Direct Diagnosis of Human Brucellosis Bacteriological Culture for Humans This is by definition the most specific diagnostic test, and since in human brucellosis it is mandatory to achieve a correct diagnosis for each individual, bacteriological cultures should be performed whenever possible. Moreover, owing to the problems posed by the serological diagnosis of B. canis (Immunological Diagnosis of Infections Caused by B. ovis and B. canis), culture is the best diagnostic tool when an infection by this species is suspected. Brucella has been isolated from several tissues, articular and cerebrospinal fluids, and bone marrow, but the culture of aseptically taken blood in tryptic soy broth or similar media (Table 10) in a 10% CO2 atmosphere is the simplest and most often used procedure. This is an enrichment procedure, and subcultures on agar media are necessary to identify the bacteria. However, blood debris prevents detection of growth by the appearance of turbidity, and unless special methods are adopted, repeated blind subculture is necessary until a successful isolation happens or the blood culture is discarded as negative. Because this procedure conveys a great risk of infection (Biosafety), Ruiz-Castañ eda developed a biphasic system for Brucella in which one of the sides of the culture flask is layered with solid medium. After inoculation, the flask is tilted every three or more days so that the broth wets the agar briefly and colonies eventually

416

E. Moreno and I. Moriyó n

appear. In addition, automated methods have been developed over the last decade that detect the CO2 resulting from aerobic bacterial growth, and they also make blind subculturing unnecessary. When using the Ruiz-Castañ eda method, blood cultures are seldom positive by the fourth day of incubation, the majority are positive between the seventh and twenty-first, and only 2% are positive after the twenty-seventh day. In this method, incubations should be carried out for at least 45 days before rejecting a blood culture as negative for Brucella (Díaz and Moriyó n, 1989). The automated CO2-detecting systems are advantageous in that they have a shorter average period of detection (usually less than five days). However, prolonged incubation is not completely avoided as some flasks do not become positive in less than three weeks. Moreover, cultures remaining negative after the first week of incubation do not always produce CO2 above the positive threshold level, and it is advisable to do blind subculturing for maximum sensitivity (Ariza, 1999; Casas et al., 1994; Yagupsky et al., 1997). The low numbers of Brucella cells in the sample (sometimes as low as 1–2 colony forming units per ml) are in all likelihood one of the reasons for prolonged incubations, and larger samples are advantageous (Zimmerman et al., 1990). It is, therefore, advisable to inoculate 5– 10 ml in duplicate flasks, as it is also advisable to perform two or three independent blood samplings at adequate intervals (Díaz and Moriyó n, 1989). Some studies have found that bacteriological detection can be improved by using the leukocyte lysis-concentration procedure (Gaviria-Ruiz and Cardona-Castro, 1995; Etemadi et al., 1984). When blood samples are taken from febrile patients, isolation is successful in the majority of cases, and much lower rates are obtained from afebrile patients (68 versus 32% in the extensive comparative study of Rodríguez-Torres et al., 1977). This difference is also found in relapsed patients. Indeed, the probability of obtaining a successful culture is greatly reduced when antibiotherapy has started. Since the percentages of positive cultures reported for B. melitensis (about 70-80%) are usually higher than for B. abortus (about 50%), it has been suggested that the infecting species influences the likelihood of a successful isolation (Dalrymple-Champneys, 1960). Unless epidemiological data are necessary, or infection with the vaccine strain US19 or Rev. 1 is suspected (Treatment), identification to genus level is enough for medical purposes (Identification and Typing). Molecular Tests for Humans A number of molecular tests have been proposed for human

CHAPTER 3.1.16

brucellosis, but PCR is the only one that has deserved significant attention. So far, amplification of DNA extracted from blood with primers taken from a 31-kDa B. abortus soluble protein (BCSP31) is the only protocol that has been evaluated (Matar et al., 1996; Morata et al., 1999a; Queipo-Ortuñ o et al., 1997). This protocol has been reported to have a diagnostic sensitivity of 92% or higher (with regard to the combined clinical, bacteriological and serological evidence) and a specificity of 93%. Modifications of the original protocol have been described in further work (Morata et al., 1998; Queipo-Ortuñ o et al., 1999), and the protocol reported to be successful in the evaluation of recovery and in the diagnosis of relapses (Morata et al., 1999b). However, other laboratories have not been able to reproduce the results (Navarro et al., 1999). If the findings are confirmed, the method would provide medical microbiologists with an exceedingly valuable diagnostic tool. Direct Diagnosis in Animal Brucellosis Bacteriological Culture for Animals Bacteriological culture, by definition, is the most specific diagnostic test in animal brucellosis, and when performed exhaustively, it allows the identification of over 90% of infected cattle, sheep and also possibly of other animal species. Unfortunately, performed in this way, it is cumbersome and expensive, and the processing of samples containing B. melitensis, B. abortus or B. suis requires special precautions (Alton et al., 1988) (Biosafety). Its use is, therefore, restricted to some circumstances, such as the unequivocal demonstration of the disease at herd level, the identification of vaccine shedders (Identification of Vaccine Strains; Epidemiology), the assessment of the performance of serological tests and vaccines, and the diagnosis of animals of high genetic value. Samples used include vaginal swabs, semen, aborted fetuses (stomach contents, lung and spleen) and fetal membranes. Direct examination of smears of these materials stained by Stamp’s method (Cellular and Colonial Characteristics) can be very informative, although other bacteria like Chlamydia are also Stamp positive and a cause of abortion (Fig. 10). Blood can also be used and hemocultures are recommended as a good diagnostic test for B. canis in dogs. Milk (cream and sediment from the four quarters) is very useful for identifying shedders. Materials that can be obtained at the abattoir or after necropsy include the udder and the uterus of pregnant or early postparturient animals and the male reproductive organs, but the best samples are the spleen and, in particular, the lymph nodes. Although mammary nodes yield Brucella

CHAPTER 3.1.16

most often, all major lymph nodes (up to seventeen) should be examined for an exhaustive inquiry into the infectious state of a particular animal. In addition, any tissues showing inflammatory lesions and abscesses should be cultured (Alton et al., 1988). Most samples require maceration (in a safety cabinet). All but blood or other normally sterile fluids (which can be processed in the same way as human blood cultures) are inoculated as abundantly as possible into an appropriate selective or enrichment medium (Table 11) and incubated in a 10% CO2 atmosphere. Cultures are seldom clearly positive by the fourth day of incubation and should be incubated for at least 15 days. The number of bacteria in the organs can vary widely and plates can contain from very few to many Brucella colonies. An alternative to selective media is the inoculation of guinea pigs followed by the recovery of the bacteria from the appropriate tissues at necropsy and the demonstration of seroconversion (Alton et al., 1988; Ne’eman and Jones, 1963; Brucellosis in Guinea Pigs). This method, however, is only justified when dealing with very heavily contaminated specimens and after consideration of the ethical issues involved. Molecular Tests for Animals Tests detecting Brucella cells or antigens by immunological methods have been described but they have not found a practical use in the diagnosis of brucellosis, and this is also true of tests detecting directly Brucella DNA with labeled probes (Mayfield et al., 1990). PCR has deserved much greater attention, but so far most research has been directed to the development of protocols under laboratory conditions (Table 4), and since studies on the actual diagnostic value are scarce, there is no established PCR protocol that could replace the bacteriological culture. An assessment of the possible practical value of the existing PCR protocols is complicated by the fact that an efficient DNA extraction from samples is critical for optimal diagnostic sensitivity. There are no specific studies on this problem, but it has been noted that even in culture, Brucella cells are resistant to conventional DNA extraction protocols (Romero and Ló pez-Goñ i, 1999). Nevertheless, the few studies that have compared the diagnostic performance of PCR with that of standard bacteriological or serological tests suggest that PCR could be a valuable diagnostic test in animal brucellosis. In the examination of tissues, organs and stomach contents of B. abortus infected cattle, a PCR protocol with primers taken from a putative B. abortus Omp showed 98% sensitivity and 96% specificity as compared to bacteriological culture (Fekete et al., 1992a; Fekete et al., 1992b). In a study carried out to

The Genus Brucella

417

assess the ability to detect Brucella in the stomach contents of aborted sheep fetuses, PCR with 16S rRNA primers performed closely to bacterial cultures, albeit the number of samples studied was low (Cetinkaya et al., 1999). For obvious reasons, milk would be one of the samples of choice, and in one study, conventional PCR with 16S rRNA primers was found to have 100% specificity and 87.5% sensitivity with respect to bacteriological culture (ELISA showed 100% specificity and 98.2% sensitivity in the same samples; Romero et al., 1995b). Other possible uses of PCR in the diagnosis of animal brucellosis include the direct identification of B. abortus strain 19 vaccine shedders by using primers taken from the ery region (Identification of Vaccine Strains). Immunological Tests for Brucellosis Brucella evokes a cell- and antibody-mediated immune response (Control of the Infection), and accordingly, immunological tests for brucellosis can be grouped into two categories: those that detect specific antibody and those that analyze cell-mediated immunity. Mainly protein antigens trigger the cell-mediated response, but also both LPSs and proteins elicit an antibody response. Thus, broadly speaking, antibody tests for brucellosis can be subclassified into two groups corresponding to each of these antigens. Although structurally related to the LPS O-polysaccharide, NH or polysaccharide B is measured by a third group of antibody tests. To understand the immunological diagnosis of brucellosis, it is necessary to keep in mind 1) the differences in surface antigens existing between smooth and rough Brucella species (Response to Environmental Stress); 2) the close structural similarity of the LPS O-polysaccharides of the Brucella serotypes; 3) the presence of the smooth LPS in the most effective vaccines (Vaccines and Vaccination); 4) the existence of Gram-negative bacteria that, albeit phylogenetically distant, show O-polysaccharides similar to those of the smooth Brucella species; and 5) that significant immunological crossreactivities with Brucella proteins have been shown only for similar antigens of phylogenetically close relatives (Fig. 3). The implications of these facts can be summarized as follows: A. LPS tests used to diagnose infections caused by Brucella rough species (B. ovis and B. canis) are different from those used to diagnose B. melitensis, B. abortus and B. suis infections. B. If quality control of antigens is always important, dissociation of smooth Brucella cultures is a constant possibility. Thus, extreme care must be taken to make sure that the antigens used in smooth LPS tests are prepared

418

E. Moreno and I. Moriyó n

from nondissociated cultures (Colonies and Dissociation). C. Antibodies to the smooth-LPS can be detected with whole smooth cells (Outer Membrane Topology) or LPS extracts. It is not necessary to use bacteria or LPS of the homologous serotypes. Thus, B. abortus biotype 1 cells (often B. abortus vaccine strain 19) can be used to obtain antigens for the diagnosis of infections caused by other smooth brucellae. This has been clearly shown for B. melitensis and B. abortus biotypes serologically different from biotype 1, and it is so because polyclonal responses contain a large proportion of antibodies that, because of combinations of titer and avidity, show overlapping reactivities with the A, M and C epitopes (Fig. 15). In the diagnosis, therefore, the C epitopes are largely dominant (Fig. 34; Response to Environmental Stress). D. Smooth LPS tests have two main sources of nonspecificity: vaccination with B. abortus strain 19 or B. melitensis Rev. 1 (Vaccines and Vaccination) and infections with Gram-negative bacteria carrying perosamine-containing Opolysaccharides. Among the latter, Y. enterocolitica O:9 is the one showing the strongest crossreactivity due to the rather close similarity existing between its O-polysaccharide and the O-polysaccharide of B. abortus biotype 1 (Table 6). E. Protein tests allow a differential diagnosis of brucellosis and infections caused by Gram-negative bacteria that carry perosaminecontaining O-polysaccharides. They are also the antigens active in tests assessing the cellmediated immunological response. Practically all types of immunological assays available have been investigated as potential tests for the diagnosis of brucellosis. The most relevant antigens and, where appropriate, immunoglobulins involved in some commonly used or representative tests are summarized in Table 16. Immunological Diagnosis of Human Brucellosis Caused by Smooth Brucella Brucella melitensis, B. abortus and B. suis are the Brucella species infecting humans most often (Brucellosis in Humans). Brucella canis is also pathogenic for humans, but it does not pose a threat to human health of the magnitude created by the first three species, and the few reported cases are not always confirmed by bacteriological cultures (Carmichael, 1990). The immunological diagnosis of B. canis infection is considered in a separate section (Immunological Diagnosis of Infections Caused by B. ovis and B. canis). Antibody Detecting Tests in Infected Patients Tests detecting cell-mediated immunity are not used nowadays for the diagnosis of human bru-

CHAPTER 3.1.16

cellosis. On the other hand, antibody-detecting tests are performed routinely and they can be performed with blood serum and, when neurobrucellosis is suspected, with cerebrospinal fluid. Since serum samples are rarely taken early enough, seroconversions are seldom observed in human brucellosis. Serological tests are then useful not only to detect anti-Brucella antibodies, but also to evaluate the stage of evolution of the disease through an assessment of the classes of immunoglobulins involved in a given test or specifically detected by it. Obviously, a dominant IgM response is characteristic of the early stages (acute brucellosis), whereas IgG and IgA in the absence of IgM are characteristic of a brucellosis that was acquired some time before the test was performed. To detect changes in the immunoglobulin levels also can be useful to evaluate the possibility that a relapse had occurred (Fig. 39). Smooth-Lipopolysaccharide Tests in Infected Patients

At the early stages of infection, antibodies are directed almost exclusively against the Opolysaccharide of the smooth-LPS, and they are also those that persist longer after recovery (Fig. 39). Thus, changes in immunoglobulin profiles are best observed in smooth-LPS tests. Moreover, these are the most sensitive tests. Agglutination tests are inexpensive, technically simple and informative. Thus, they are recommended for the diagnosis of human brucellosis in most situations. They include the classical serum tube agglutination test used to titrate the total amount of agglutinating antibody and modified tests aimed to roughly determine the amount of IgM versus IgG plus IgA by including reagents that preferentially (but not exclusively) inactivate IgM (e.g., the β-mercaptoethanol test; Table 16). The diagnostic titer in the classical tube agglutination test may vary depending upon the prevalence and other circumstances (Díaz and Moriyó n, 1989). Agglutination tests for brucellosis also include the rose bengal test (and the card test), in which the use of an acid pH makes all antibodies show their intrinsic agglutinating activity. Although it is primarily a rapid slide agglutination screening test, the rose bengal can also be used in human brucellosis with serum dilutions to obtain a rough idea of the total amount of anti-smooth-LPS antibody, which is enough for most clinical purposes. Prozones are occasionally observed in the classical tube agglutination, but not in the rose bengal, and both tests have very few falsenegative results. The serum agglutination test can be adapted to measure the nonagglutinating (low affinity) IgG and IgA (Coomb’s test; Table 16), which characteristically develop in brucellosis. In acute brucellosis, Coomb’s titers are usually 4–16 times

Precipitation Double immunodiffusion

Complement fixation

Complement consumption Hemolysis in agar

Extracts containing S-LPS, S-LPS plus NH, or R-LPS plus Omps (B. ovis hot saline extracts)

S-LPS (B. abortus cell suspension) or R-LPS plus Omps (B. ovis hot saline extracts)

S-LPS coated erythrocytes

S-LPS (stained B. abortus cell suspension in citrate buffer pH 4.0)

Milk ring test

Coomb’s

S-LPS (stained B. abortus cell suspension in lactate buffer pH 3.65) S-LPS (B. abortus cell suspension)

S-LPS (B. abortus cell suspension)

Antigen

Rose bengal (and card) tests

Agglutination Tube agglutination (and β-mercaptoethanol and rivanol) test

Tests

Table 16. Some tests and antigens used in the diagnosis of brucellosis.

IgG (also IgM)e

IgGd

IgGd

IgA (also IgM and IgG)

IgG and IgA

IgM, IgG and IgAc

IgM, IgG and IgA

Antibody

Used mostly for B. ovis and B. canis diagnosis with the R-LPS plus Omps extract; usually performed in hypertonic gels

Sensitized erythrocytes are included with complement in an agarose gel and a radial immunodiffusion performed A confirmatory test for B. abortus or B. melitensis animal brucellosis; also used in the diagnosis of B. ovis. Cumbersome to perform unless automated; some sera show anticomplementary activity

It can be adapted to the diagnosis of B. canis. To estimate the amount of IgM, the results of the standard tests are compared with those obtained in the presence of β-mercaptoethanol (or similar reagents)a or rivanolb Simple and inexpensive slide-agglutination type tests in which bacteria are stained with rose bengal. Used as screening tests Detects nonagglutinating Ig that has reacted with the bacterial surface by using an anti-Ig serum that brings about the agglutination. It can, therefore, be used to specifically detect nonagglutinating IgG or IgA with an antiserum of the appropriate specificity Ig in milk reacts with hematoxylin stained bacteria, and the complex attaches to the fat that ascends to form a ring on the surface of the reaction tube. Only adequate for cow milk

Comments

Alton et al., 1988

Alton et al., 1988

Ruckerbauer et al., 1984

Alton et al., 1988 Sutra et al., 1986

Otero et al., 1982

Alton et al., 1988

Alton et al., 1988

References

CHAPTER 3.1.16 The Genus Brucella 419

Soluble proteins of rough Brucella mutants

Complex antigenic mixture; soluble and outer mebrane proteins Soluble proteins of rough Brucella mutants

Delayed type hypersensitivity

Lymphocyte blastogenesis

Antibody

None

None

None

Presumably all Ig classes

Presumably all Ig classes

Depends on conjugate specificity

IgG

IgG

f

Technically demanding

Technically demanding

A mouse monoclonal antibody competes with low avidity (and low titer) antibodies for the S-LPS C epitopes; the monoclonal antibody bound is then detected with an immunoconjugate specific for mouse immunoglobulins A promising test based on the change of fluorescence of labeled antigen upon antibody binding Most often performed as a simple skin test

In human brucellosis, ELISAs detecting anti-S-LPS antibodies could replace classical tests if adapted to measure IgM and IgG. For animal brucellosis, ELISAs are most often adapted to detect anti-S-LPS IgG

A confirmatory test for cattle, goat and sheep (B. melitensis) brucellosis. Performed in hypertonic gels Useful for the diagnosis of human brucellosis. IgM to protein antigens is not detected in this assay

Comments

Weynants et al., 1995

Jones et al., 1973 Blasco et al., 1994 Saegerman et al., 1999 Baldwin et al., 1984

Nielsen et al., 1996

Nielsen et al., 1996 Alonso-Urmeneta et al., 1988 Marín et al., 1998 Rosetti et al., 1996 Goldbaun et al., 1992 Letesson et al., 1997 Gall and Nielsen, 1994

Díaz et al., 1976

Jones et al., 1980 Díaz et al., 1981

References

E. Moreno and I. Moriyó n

b

a

At least human IgA has been shown to be sensitive to the concentrations of β-mercaptoethanol used in the test. 2-Ethoxy-6,9-diamino-acridine (an IgM precipitating reagent). c IgA activity in the rose bengal test has been shown with human sera. d There are contradictory reports on the IgM ability to activate complement. At least in cattle, IgM does not activate guinea pig complement in the absence of homologous serum components. e IgM is particularly active in the immunoprecipitation of LPS extracts (this molecule forms aggregates, and the reactions mimic a microagglutination). f IgM precipitating NH has been shown in some human sera. At least in cattle, IgG1 is the major immunoglobulin precipitating NH (and polysaccharide B) and accounting for the hypertonic conditions required in the assay.

γ-interferon production

Fluorescein-labelled Opolysaccharide

S-LPS, S-LPS O-polysaccharide, or S-LPS rich extracts

Fluorescence polarization

Competitive ELISA

Purified S-LPS, S-LPS Opolysaccharide, S-LPS rich extracts or extracts containing R-LPS plus Omps (B. ovis hot saline extracts). Purified proteins

Soluble proteins of rough Brucella mutants

Crossover immunoelectrophoresis

Immunoenzymatic Indirect ELISA

NH and polysaccharide B

Antigen

Reverse radial immunodiffusion (RID)

Tests

Table 16. Continued

420 CHAPTER 3.1.16

CHAPTER 3.1.16

higher than tube agglutination titers, whereas at later stages they are 16–256 times higher. Thus, comparison of the serum agglutination and the Coomb’s tests allows the assessment of the stage of evolution of human brucellosis. This can also be achieved by using the complement fixation test: in over 90% of the human cases, serum agglutination and complement fixation are simultaneously positive, but after the fourth or the fifth week of evolution, complement fixation titers are higher than serum agglutination titers. At the very early stages of infection, serum agglutination is positive and complement fixation negative, and the opposite is true in patients that are recovered or are developing chronic or focal forms (Díaz and Moriyó n, 1989). The indirect enzyme-linked immunosorbent assay (i-ELISA) with smooth-LPS (or with smooth cells) is both very specific and sensitive for the diagnosis of human brucellosis. Using this assay, it has been clearly proven that the response to smooth-LPS in treated patients follows a classical pattern: IgM appears and decreases faster than IgA or IgG, and in relapses, IgG and IgA, but not IgM, increase over the levels observed when the first diagnosis was made (Fig. 39). The assay also shows that IgG is the more persisting immunoglobulin and that over 80% of the patients have high titers of IgG and IgA a year and a half after clinical recovery (Ariza et al., 1992; Pellicer et al., 1988). The correlation of the IgM or IgG i-ELISA with agglutination, Coomb’s and complement fixation tests is the one expected from the immunoglobulin classes involved in the latter tests. It seems, therefore, possible to replace all classical serological tests by an i-ELISA with smooth-LPS and anti-IgM and IgG conjugates, but this would require not only standardization but also the definition of the equivalence between the i-ELISA results and those of the classical tests that clinicians are used to interpreting. There is one report on the use of the competitive ELISA (c-ELISA) in human brucellosis, and the assay was found also very specific and sensitive (Lucero et al., 1999). This test could be less sensitive to interference by crossreacting bacteria, but no definite studies with human sera have been published to date. Protein Tests in Infected Patients Both indirect evi-

dence and studies in experimentally infected animals show that antibodies to soluble proteins develop later than antibodies to the smooth-LPS. They are characteristic of an active brucellosis, and the longer the evolution of the disease before treatment, the higher the antibody levels and the diversity of proteins to which they are directed (Fig. 38). Antibodies to Brucella proteins can be found not only in blood serum but

The Genus Brucella

421

also in the cerebrospinal fluid of patients afflicted with neurobrucellosis. The protein antigens that have been used include the mixture of soluble proteins present in the hypertonic Brucella extracts also used in the skin test (Díaz et al., 1976; Cell-mediated Immunity Tests), BP26 (Rossetti et al., 1996) and an immunocaptured cytosolic protein (lumazine synthase; Goldbaum et al., 1992; Goldbaum et al., 1999). Antibodies to Omps also develop in brucellosis but they are more cumbersome to detect and their diagnostic value is less clear (Leiva-Leó n et al., 1991). Tests that have been used include gel precipitation (either as double gel diffusion or as crossover immunoelectrophoresis), indirect ELISA and Western blot. By gel immunodiffusion, it has been shown that over 80% of brucellosis patients have precipitating antibodies to one or several soluble proteins in hypertonic Brucella extracts. In one study, in which the sera of 173 patients with acute brucellosis and 39 with brucellosis having a long evolution were examined by crossover immunoelectrophoresis, 84.9% and 91.6% had developed anti-protein antibodies, respectively (Díaz et al., 1976). Using an indirect ELISA in which the antigen had been immunocaptured with a monoclonal antibody, 94% (33 of 35) of the brucellosis patients examined showed IgG to Brucella lumazine synthase (an 18-kDa antigen), as well as strong anti-smooth-LPS response. By contrast, individuals classified as having a “nonactive” brucellosis seldom showed anti-lumazine synthase antibody, even though most had various degrees of anti-LPS reactivity (Goldbaum et al., 1992). All these results strongly suggest that the presence of anti-protein antibodies is a useful criterion to decide whether low levels of antismooth-LPS antibodies are indicative of infection or simply of contact without the development of disease. Thus, these tests deserve further attention. Antibody Detecting Tests in Recovered Patients After recovery, serum agglutination becomes progressively negative, usually six months to two years later. The complement fixation titers also decrease but they may also persist, always for longer times than those of serum agglutination titers. The last conventional test to become negative is the Coomb’s test. When the results of these three classical tests are taken together, the serology seldom becomes negative by the third month, and positive results often persist after the eighth month of evolution. In the ELISA, IgM, IgA and IgG are negative by the first year in 65, 25 and 3% of recovered patients, respectively (Ariza et al., 1992). The problem of predicting a relapse based on the results of serological tests has not been satisfac-

422

E. Moreno and I. Moriyó n

torily solved. It has been suggested that persisting β-mercaptoethanol-resistant antibodies (IgG) correlate with the risk of relapse, but the ELISA studies show that IgG persists for a long time in the serum of fully recovered patients. Immunological Diagnosis of Animal Brucellosis Caused by Smooth Brucella A major difference with the immunological diagnosis in humans is that, in animals, what is most often judged is whether there has been contact with Brucella and not the existence of an ongoing infection. This information is enough for eradication purposes, including control of animal movements and trade, and an assessment of the immunoglobulin profile or a comparison of the results of smooth-LPS and protein tests is not necessary. However, the use of vaccines complicates the diagnosis of animal brucellosis. Vaccination induces an immunological response that closely mimics that resulting from a true infection, and since even the best vaccines available do not afford 100% protection, distinction of infected and vaccinated animals becomes necessary. For obvious reasons, the majority of the information on the performance of the immunological tests concerns cattle, sheep, goats and swine. Positive reactions in the classical blood serum tests have been recorded for other important domestic animals, such as camels, water buffalos and reindeer, and also for a variety of wild animals (Davis, 1990). However, since these observations have not been complemented by bacteriological studies, the actual performance of the tests in detecting Brucella infection in all these animals is not known and the results of the immunological tests can only be taken as presumptive. The discussion is limited here to the immunological diagnosis of brucellosis in domestic ruminants and swine. Antibody Detecting Tests in Infected Animals The tests used are often the same as those used in human brucellosis (Table 16), and in fact, some, like the rose bengal test, were first developed for the diagnosis of animal brucellosis. The presence of antibodies to Brucella can be investigated in blood serum but also in the whole milk or whey of dairy animals. Smooth-Lipopolysaccharide Tests in Infected Animals Like in human brucellosis, these are the

tests used most often. Their application has been studied best in cattle. There are fewer studies in small ruminants, and the number of animals included is also comparatively reduced. Moreover, for classical tests, such as the rose bengal and complement fixation, most investigations in small ruminants have been performed with the tests standardized for the diagnosis in cattle, and

CHAPTER 3.1.16

these conditions do not result in optimal sensitivity (Blasco et al., 1994b). This standardization problem affects only the proportion of serum to antigen and not to the particular epitopic structure of the smooth-LPS used in the assay (Immunological Tests for Brucellosis). The results of selected studies in which this difficulty is taken into account, the sensitivity defined with respect to bacteriological culture, and the specificity determined in the absence of vaccination are presented in Table 17. They show that, under those circumstances, most tests perform with a remarkable high efficiency and that the i-ELISA and the rose bengal (or card) test come close to the ideal test. There is a long practical experience in the use of the rose bengal test, but the iELISA has been investigated thoroughly. It has been shown that purified smooth LPS, Opolysaccharide or NH preparations offer no advantage over relatively crude smooth-LPS extracts (Alonso-Urmeneta et al., 1988; Nielsen et al., 1995; Wright and Nielsen, 1990b). Conjugates based on monoclonal antibodies specific for ruminant IgG1 or on the IgG-binding Streptococcus protein G allow the diagnosis of the three common domestic ruminants with a single reagent, improving the specificity and reducing the background reactivity caused by IgM (Alonso-Urmeneta et al., 1988; Wright et al., 1990b). Peroxidase conjugates and the chromogenic substrate 2,2′-azino-bis(3ethylbenzthiazoline-6-sulfonic acid) (ABTS) have been shown to produce excellent results. Thus, this assay has been well-defined and the choice between the i-ELISA and the rose bengal test depends mostly on economical and logistic reasons. With regard to swine brucellosis, studies with well-defined sera are scarce but immunoenzymatic and fluorescence polarization assays seem to outperform the classical agglutination or complement fixation tests (Table 17). The latter tests have been reported to show low sensitivity and to be useful for diagnosis only on a herd basis (Alton, 1990a). The specificity of the smooth-LPS tests becomes compromised when vaccination with B. abortus 19 or B. melitensis Rev. 1 is implemented (Vaccines and Vaccination). To overcome this problem (and also to reduce the restrictions in the application of the vaccines), a test that is able to distinguish infected from vaccinated animals (i.e., a test showing a high specificity in the context of vaccination) would be necessary. Ideally, such a test should also have a high sensitivity and be technically simple to facilitate large-scale testing. No classical test fulfils these requirements, and therefore a combination of a simple screening test (highly sensitive) and a confirmatory test (highly specific with respect to vaccinated animals) has been traditionally used. The rose ben-

CHAPTER 3.1.16

The Genus Brucella

423

Table 17. Performance of some antibody tests for the diagnosis of infections by smooth Brucella spp.

Animal host Cattle

Test

Sensitivitya (%)

Specificityb (%)

No. of culturepositive animals studied

Rose bengal (or card-test)

91.4–99.7

98.8–99

300 and 1,666

87.9 or 97.1c 92

93.1 or 9.8c 100

654 112 and 383

i-ELISA-LPS c-ELISA-LPS Fluorescence polarization Rose bengal Complement fixation RID-NH

100 100 99 94 88 90–96

99.7 99.9 99.9 100 100 100

636 636 561 135 135 85 and 77

i-ELISA-LPS

100

100

55 and 140

Complement fixation RID-NH

Sheep

Goats

Swine

c-ELISA-LPS Rose bengal Complement fixation RID-NH i-ELISA BPATd Complement fixation i-ELISA c-ELISA Fluorescence polarization

96 100 94 94 100 77 95.5–99.9c 94 96 93.5

100 100 100 100 100 95.9 58.1–93.2c 98 90 97.2

55 55 55 55 55 401 401 401 401 401

References Huber and Nicoletti, 1986 Morgan, 1977 Nielsen et al., 1996b Díaz-Aparicio et al., 1993 Jones et al., 1980 Nielsen et al., 1996b Nielsen et al., 1996b Nielsen et al., 1996a Blasco et al., 1994a Blasco et al., 1994a Díaz-Aparicio et al., 1993 Jimenez Bagués et al., 1992 Blasco et al., 1994b Marin et al., 1994 Marin et al., 1999 Díaz-Aparicio et al., 1994 Díaz-Aparicio et al., 1994 Díaz-Aparicio et al., 1994 Díaz-Aparicio et al., 1994 Nielsen et al., 1999 Nielsen et al., 1999 Nielsen et al., 1999 Nielsen et al., 1999 Nielsen et al., 1999

Abbreviations: RID-NH, radial immunodiffusion for antibody to native hapten polysaccharide; i- and c- ELISA, indirect and competitive enzyme-linked immunosorbent assay; and BPAT, buffered antigen plate agglutination test. a Sensitivity = number of test-positive sera among culture-positive animals / total number of culture positive animals. b Specificity = number of test-negative sera among Brucella-free (unvaccinated) animals / total number of Brucella-free (unvaccinated) animals. c Depending on whether sera with anticomplementary activity are considered as positive or negative. d BPAT, buffered plate agglutination test. This is a rapid agglutination test similar to the rose bengal test, but cells are stained with a mixture of crystal violet plus brilliant green. It may be slightly more sensitive than the rose bengal test.

gal and the complement fixation, respectively, have been the tests of choice for these purposes. The former test is simple, can be automated, and is very sensitive (Table 17), but it remains positive in a large proportion of vaccinated animals long after vaccination. The complement fixation test yields negative or weak titers six months after calfhood vaccination, but it requires careful and periodical standardization, and when adult vaccination is used, some animals may have titers persisting for a long time. Despite these problems, the efficacy of the combination of these two tests with calfhood vaccination is well established (MacMillan, 1990a), and they have been the reference in the serological diagnosis of animal brucellosis for years. Although it has been accepted that their performance in sheep and goats is adequate, they lack sensitivity when standardized according to current guidelines (Blasco et al., 1994b).

Several studies have shown that the i-ELISA is highly sensitive and, when properly standardized, also specific in differentiating infected from vaccinated animals, with reported specificities as high as 95–98% for cattle (Samartino et al., 1999). In experiments performed in sheep bled periodically after vaccination, the i-ELISA was less specific than the complement fixation, an immunoprecipitation test with NH (NH [and Polysaccharide B] Tests), and a competitive ELISA (c-ELISA; Marín et al., 1999). This last type of assay has been designed to prevent reactions below a given avidity threshold level set by the concentration of a competing monoclonal antibody specific for the O-polysaccharide C epitope(s) (Table 16). Since there is evidence that the antibodies remaining in serum several months after vaccination are in most cases of low avidity, the c-ELISA is an interesting approach to the differential diagnosis of infected and vac-

424

E. Moreno and I. Moriyó n

cinated animals. An additional advantage of this method is that it does not use a conjugate specific for the immunoglobulins in the tested serum, and therefore, it can be easily adapted to detect Brucella infections in different animal species. Like the i-ELISA, several protocols are possible with regard to antigen, monoclonal antibody, and other conditions. In a study performed in cattle, the sensitivity of a c-ELISA with monoclonal BM-40 was 100% in animals with bacteriologically proven infection, but its specificity in vaccinated calves was only 80% (76% for the complement fixation in the same animals; Macmillan et al., 1990b). However, an improved c-ELISA with monoclonal M48 showed 100% specificity in differentiating infected from vaccinated cattle (Nielsen et al., 1995). In a large study conducted to evaluate several i-ELISA and cELISA protocols with sera from animals of unknown bacteriological and vaccine status, the c-ELISA with smooth LPS and monoclonal M48 was found to be the best performing assay in terms of balanced sensitivity and specificity (97.5 and 96.5%, respectively) with respect to the combined results of rose bengal and complement fixation (Gall et al., 1998). In the analysis of sera from Brucella-free, vaccinated, and bacteriologically positive sheep, this same c-ELISA showed slightly lower sensitivity than the i-ELISA (96 and 100%, respectively) and better specificity (about 89%) in distinguishing infected from Rev. 1 vaccinated animals (Marín et al., 1999). In this same study, the c-ELISA was more specific (87– 89%) than the complement fixation in vaccinated adults and in subcutaneously vaccinated young sheep (45–66 and 78%, respectively) but not in conjunctivally vaccinated young sheep (89% for the c-ELISA versus 100% for the complement fixation). Thus, when properly standardized, the c-ELISA seems a better test than the complement fixation but the influence of the route of vaccination deserves further research. NH (and Polysaccharide B) Tests Immunoprecipitation tests with NH or polysaccharide B are very specific when cattle are vaccinated with B. abortus strain 19 and can replace the complement fixation test. For optimal sensitivity (up to 92%), precipitating antibodies are detected in a reverse radial immunodiffusion (RID) system, in which the serum is allowed to diffuse in a hypertonic gel containing the polysaccharide (Díaz et al., 1979; Díaz et al., 1981), but the more simple double gel diffusion procedure is also useful (Lord et al., 1989; Pinochet et al., 1989; Fig. 19). Calves vaccinated at 3–5 months of age are negative after 2 months of vaccination, and adult cattle vaccinated 4–5 months previously with the standard dose of B. abortus 19 do not give positive reactions unless the animals develop vaccine

CHAPTER 3.1.16

infections in the mammary gland and shed the vaccine in their milk (Jones et al., 1980). In a study carried out with the sera of 1,057 cattle from infected herds in which adult vaccination was implemented, the RID test outperformed the complement fixation in specificity (80 versus 66%, respectively) and showed acceptable sensitivity (88 versus 98%, respectively) despite the fact that vaccination and infection were superimposed (Jones et al., 1980). The RID test also shows better specificity than the complement fixation test in adult and young sheep vaccinated with Rev. 1, either subcutaneously or conjunctivally (Blasco et al., 1984a; Jiménez de Bagués et al., 1992). A remarkable characteristic of the RID test is that a positive result correlates with Brucella shedding. In addition to the evidence mentioned above on the strain 19 shedders, this has been shown in experimentally infected cattle (Jones et al., 1980) and also in naturally infected cattle undergoing antibiotic treatment (Jiménez de Bagués et al., 1991). Thus, the combined use of rose bengal, RID and strain 19 vaccination has been proven to be useful in achieving eradication (Asarta, 1989). The NH (and polysaccharide B) and the LPS O-polysaccharides are structurally very similar (Fig. 16; Response to Environmental Stress). It is thus noteworthy that whereas both antigens yield very close results when used in the iELISA, immunoprecipitation with smooth LPS has a much lower sensitivity than immunoprecipitation with NH (Alonso-Urmeneta et al., 1988). These differences in sensitivity can be explained by the fact that the high epitopic density and disperse state of NH (versus the bulky micellelike state of LPS) favor its immunoprecipitation when only low titers of antibody are present. Moreover, the comparatively high threshold antibody avidity required for a positive immunoprecipitation could account for the ability of the RID with NH to discriminate infected from vaccinated animals, since it has been observed that unless udder infections develop, vaccination only transiently induces the high avidity IgG that results from an active infection (B. AlonsoUrmeneta et al., unpublished observations). Comparative studies between immunoprecipitation with NH and c-ELISA show similar sensitivity (90 and 96%, respectively) and specificity in vaccinated animals (85–100% and 87–98%, respectively; Marín et al., 1999), supporting by this manner the interpretation that the former assay detects only high avidity antibodies. It seems likely that the subtle structural and epitopic differences existing between the NH (and polysaccharide B) and the O-polysaccharide could contribute to the peculiar immunoprecipitating properties of the former polysaccharide. In one study in which it was com-

CHAPTER 3.1.16

pared with B. melitensis 115 polysaccharide B, the O-polysaccharide obtained directly by hydrolysis of the smooth LPS of B. abortus strain 19 (which is poor in NH) did not produce good results (Jones et al., 1980). Although the polysaccharides obtained by acid hydrolysis of smooth virulent B. abortus produce satisfactory results (Cherwonogrodzky and Nielsen, 1988), these whole cell extracts contain both NH and Opolysaccharide, and it is not possible to determine to what extent each of these two molecules accounts for the results. Protein Tests in Infected Animals Tests detecting

antibodies to Brucella proteins are receiving an increased attention in the hope of improving the differential diagnosis of infected and vaccinated animals. If sensitive enough, these tests would also be very useful to avoid false positive results caused by bacteria crossreacting with smooth Brucella at the LPS level. Early work carried out by crossover immunoelectrophoresis showed that extracts containing a mixture of soluble proteins were promising diagnostic antigens in sheep (Trap and Gaumont, 1982). On the other hand, a gel immunodiffusion test with a purified protein (protein α-2) showed only low sensitivity (55%), even though it was highly specific (99%) in differentiating infected from vaccinated cattle (Dubray, 1984). More recently, several cloned proteins have been tested in immunoenzymatic assays. Although cattle, goats and sheep infected by smooth Brucella develop antibodies to group 1 (89 kDa), Omp2 (porins), Omp3a (also known as Omp25), Omp19, Omp16 and Omp10, the response is of lower intensity and frequency than that to the smooth-LPS, and these Omps used individually would have no diagnostic use (Cloeckaert et al., 1992; Letesson et al., 1997; Zygmunt et al., 1994). Cytosolic or periplasmic proteins tested include GroEL (Lin et al., 1996), Cu++-Zn++ superoxide dismutase (Tabatabai and Hennager, 1994), and a variety of soluble proteins of 9, 15, 17, 24, 28, 32, 39, 50 and 54 kDa (Letesson et al., 1997; Salih-Alj Debbarh et al., 1995), but lumazine synthase (an 18-kDa protein; Baldi et al., 1996) and BP26 (Arese et al., 1999; Rossetti et al., 1996; Salih-Alj Debbarh et al., 1996) seem to be the most promising reagents. It seems likely that for maximal sensitivity, several of these proteins would have to be combined in a single immunoenzymatic test. Milk Tests In dairy cattle, milk and milk serum are suitable samples either for individual diagnosis or for monitoring the status of the herd by checking pooled milk. For these purposes, the milk ring test (Table 16) was originally developed. This test uses whole milk rather than

The Genus Brucella

425

serum, has been standardized and is highly sensitive and easy to perform. Moreover, the conditions of use for testing pooled milk (i.e., sensitivity with regard to numbers of animals and total volume of sample) have been well studied (Pietz, 1977). Unfortunately, as the test depends on characteristics such as the content and type of fat in the milk, it is not applicable to small ruminants. Several i-ELISA protocols have been evaluated as alternatives to the milk-ring test in both cattle and small ruminants. Most authors have concluded that it is an excellent test for the examination of milk from individual animals (Bercovich and Taaijke, 1990; Heck et al., 1980; Mikolon et al., 1998; Romero et al., 1995b; Thoen et al., 1979; Vanzini et al., 1998). In addition, some studies have addressed the problem of determining the conditions under which the i-ELISA could be used to detect anti-Brucella antibody in bulk milk. In the analysis of a large number of farms, an i-ELISA with smooth-LPS showed better sensitivity than the milk-ring test when both were applied to bulk milk (Kerkhofs et al., 1990), and the sensitivity of the assay can be improved by concentrating the antibody in the whey (Forschner et al., 1989). Poorer results have been reported by some authors possibly owing to methodological differences, but it can be concluded that when properly standardized, the i-ELISA is a suitable method for monitoring the status of herds in Brucella-free areas (Knosel et al., 1991). Cell-Mediated Immunity Tests The value of the demonstration of a delayed-type hypersensitivity response for the diagnosis of animal brucellosis has been analyzed repeatedly. The test is performed by injection of a suitable protein allergen into the skin, and the demonstration of a characteristic lesion 48–72 hours later is taken as a positive result. LPS-free antigens are necessary because the O-polysaccharide induces an Arthustype reaction that obscures the interpretation of the test, and the toxicity of large quantities of the LPS lipid A also induces a local reaction, which obscures (without histological examination) a true delayed-type hypersensitivity response. Most often, the antigen used is a mixture of proteins extracted with NaCl from rough B. melitensis 115 (brucellin or brucellergene; Bhongbhibhat et al., 1970; Jones et al., 1973), and the use of the whole cytosolic fraction or of purified proteins does not offer advantages (Blasco et al., 1994a) nor seems to improve the sensitivity (Denoel et al., 1997b). It is accepted that the skin test is less sensitive than the serological tests, but also that it may be positive in infected animals that have serologically negative responses (Nicoletti and Winter, 1990b). In a recent study per-

426

E. Moreno and I. Moriyó n

formed in experimentally infected cattle, a sensitivity of 78–93% was reported depending on the time lapse after infection (Saegerman et al., 1999). In a study in naturally infected sheep, the skin test showed 97% sensitivity with respect to the findings of a thorough bacteriological search, but it was also positive in many animals from which Brucella was not isolated (Blasco et al., 1994a). Thus, although it seems that the skin test is not reliable for detecting individually infected animals, it may be useful as a herd test. Its use does not resolve the problem of the differentiation of infected and vaccinated cattle. Other tests that correlate with cell-mediated immunity and that have been evaluated for diagnostic purposes are the lymphocyte stimulation assays and the determination of γ-interferon (Interleukins in Brucellosis). The former tests have not been successful in distinguishing infected from vaccinated animals (Nicoletti and Winter, 1990b), and as they are expensive and technically demanding, they are not used presently. The γ-interferon assay has been evaluated preliminarily in experimentally and naturally infected heifers (Weynants et al., 1995) and reported to be more sensitive than any combination of serological tests, but its specificity has been questioned (Kittelberger et al., 1997). Immunological Diagnosis of Infections Caused by B. ovis and B. canis The main differences between the tests used in the diagnosis of infections by smooth Brucella and B. canis and B. ovis relate to their markedly different LPS structure (Response to Environmental Stress) and outermembrane topology. Because rough Brucella cells do not form stable suspensions and are prone to autoaggregate, agglutination tests produce a high proportion of false-positive results. Although this problem can be overcome in a variety of ways, the absence of the O-polysaccharide in the rough Brucella spp. comparatively reduces the importance of the antibody response to the LPS and increases that to other surface antigens. This has been clearly shown in B. ovis ram epididymitis where the antibody response to Omps, in particular to those of group 3 (Outer Membrane Proteins), is more intense than in the infections caused by B. melitensis (Riezu-Boj et al., 1990). However, neither the LPS nor the Omps by themselves are effective diagnostic antigens (Ficapal et al., 1995; Kittelberger et al., 1998; Riezu-Boj et al., 1986). This explains why serological tests for rough Brucella spp. are most effective when the hot-saline extract (Myers and Siniuk, 1970) is used because it is enriched in both rough LPS and group 3 Omps (Riezu-Boj et al., 1990). A large variety of assays have been proposed for the diagnosis of B. ovis sheep brucellosis

CHAPTER 3.1.16

(reviewed by Blasco, 1990). However, only the complement fixation test, a gel immunoprecipitation assay, and some i-ELISAs have been extensively used or tested. Brucella ovis antigens often show anticomplementary activity, and although the effect is reduced when the hot saline or similar extracts are used, it is still significant. A sensitivity of about 90% has been estimated for the complement fixation test, similar to that found for the gel immunoprecipitation with the same antigen. The i-ELISA has been studied with respect to the antigen (purified rough LPS, hot saline extract, cytosoluble proteins) and conjugate (polyclonal of heavy and light chain IgG specificity, anti-ruminant IgG monoclonal and protein G; Ficapal et al., 1995; Marín et al., 1989b, 1998; Vigliocco et al., 1997). In addition to the demonstration that hot saline extract is the best antigen in the i-ELISA, these studies have shown that the monoclonal or protein-G based conjugates reduce the background reactivity observed with the sera of healthy animals. The sensitivity of the i-ELISA has been found to be superior (Cho and Niilo, 1987; Spencer and Burgess, 1984) or equal to that of the gel immunoprecipitation or complement fixation, and there are no objective reasons to use these methods instead of the classical tests. However, since hot saline extracts are obtained in low yields and the i-ELISA uses much less antigen, it could be more practical. Brucella melitensis Rev. 1 is used to vaccinate sheep against both B. melitensis and B. ovis (Vaccines and Vaccination), and it elicits a strong antibody response to the O-polysaccharide, interfering in the smooth LPS tests. Moreover, it also elicits antibodies to the core and lipid A sections of the LPS, and these antibodies interfere in the serological tests for the diagnosis of B. ovis, owing to the shared epitopes of the inner LPS sections of B. melitensis and B. ovis (The Lipooligosaccharide of Rough Brucella Species). This interference is reduced in the case of the complement fixation and gel immunoprecipitation tests (Blasco, 1990) and more importantly in the iELISA, and this is probably due to the favorable exposure of the common epitopes after adsorption to polystyrene (Marín et al., 1998). The immunological diagnosis of B. canis infections has been less investigated, and existing tests may provide a large proportion of false-negative and false-positive results. In experimentally infected dogs, blood cultures were positive several weeks before a positive result was obtained in any of the agglutination and gel immunoprecipitation tests available (Carmichael et al., 1984). False-positive results relate to the nature of the antigens, since the preparation of bacterial suspensions efficient in agglutination tests is even more difficult than in the case of B. ovis.

CHAPTER 3.1.16

Brucella canis almost always produces mucoid colonies on standard media and bacteria strongly autoaggregate at pH below neutrality, a problem also existing with the hot saline extracts obtained from mucoid bacteria (Carmichael, 1990). The problem is partially solved using growth media and buffers at pH 7.4, and B. ovis and a nonmucoid B. canis variant have been used in an attempt to develop suitable agglutination tests (Carmichael, 1990). Moreover, it has been shown that a brief treatment with β-mercaptoethanol reduces the number of false-positive results. However, even under the best conditions, the demonstration of a specific immunological agglutination using B. ovis and B. canis cell suspensions is not obvious and requires some training (Carmichael and Shin, 1996). Agglutination can be examined in tubes by the serum dilution method or in a rapid slide test, and the interpretation has to take into account the possibility (25–50%) of a false-positive result. On the other hand, the test is accurate in identifying noninfected dogs when a negative result persists after several bleedings. As an alternative to agglutination tests, gel immunoprecipitation has been investigated. When used in this assay, hot saline or sodium deoxycholate extracts suffer from the same problems of nonspecific reactions as do agglutination tests. On the other hand, cytoplasmic protein antigens do not produce false— negative results and the method is considered to be highly accurate in detecting antibodies. Antibodies to proteins develop late after infection, and the sensitivity is low during the first months. However, these antibodies persist in chronically infected dogs at times when the other serological tests may have become negative (Carmichael et al., 1984). Immunological Diagnosis of Brucellosis and Cross-Reacting Bacteria Smooth-LPS tests are susceptible to yield false-positive results when infections by Gram-negative bacteria with perosamine-containing O-polysaccharides occur (Table 6). But for those due to Y. enterocolitica O:9, crossreactivities are mild, and none is a significant problem in the diagnosis of human brucellosis when the clinical symptoms and the results of the anamnesis are considered. Indeed, in addition to some of the serological tests discussed in the following paragraphs, blood cultures are 100% specific. On the other hand, Y. enterocolitica O:9 infections in cattle can be troublesome, particularly in countries free from B. abortus, since the sporadic appearance of positive serological results among unvaccinated populations calls for an immediate differential diagnosis. Although orally acquired Y. enterocolitica O:9 seldom induces high levels of anti-

The Genus Brucella

427

bodies to smooth Brucella LPS and the responses are transient in cattle (Garin-Bastuji et al., 1999; Kittelberger et al., 1997; Mittal et al., 1985), titers in blood serum and milk may be higher and persistent when mammary infections occur (Mittal et al., 1985). The problem can also be a significant one in swine (Wrathall et al., 1993). So far, three different approaches have been followed to develop differential tests. The demonstration of antibodies specific for the enterobacterial common antigen (Mittal et al., 1980b), Y. enterocolitica flagellar antigens (Mittal and Tizard, 1979) and Yersinia outer proteins (yops; Kittelberger et al., 1995b; Weynants et al., 1996a; Weynants et al., 1996b; Yersinia) has been proposed to identify with precision those cases suspected of Y. enterocolitica O:9 infection. However, antibodies to antigens shared by enteric bacteria are not uncommon in healthy cattle and titers of anti-flagella antibody are weak possibly because Y. enterocolitica is nonmotile at the host body temperature. Moreover, dual infections by Yersinia and Brucella do not seem uncommon in livestock, and this reduces the usefulness of the yops (Kittelberger et al., 1995b; Mittal and Tizard, 1980a). On the other hand, yops have been found clearly useful for the differential diagnosis in humans (Schoerner et al., 1990). An alternative approach is based on the small differences existing between the LPS of Y. enterocolitica O:9 and smooth Brucella spp. at Opolysaccharide (Response to Environmental Stress) and core oligosaccharide levels. Early reports suggest that whereas the antigenic determinants involved in the crossreaction are resistant to periodate oxidation and borohydride reduction, the determinants specific for Y. enterocolitica O:9 LPS are sensitive and on this basis an i-ELISA with both treated and untreated LPS was proposed for the differential diagnosis of human infections (Lindberg et al., 1982). The differences at LPS level were also noted in competitive immunoassays (Granfors et al., 1981). Based on the results of experimental infections in cattle, the c-ELISA has been proposed as a specific diagnostic test that is able to discriminate antibody elicited by Y. enterocolitica O:9 (Nielsen, 1990a). A c-ELISA with a monoclonal antibody to the C/B epitope (shared by smooth Brucella spp. and absent from Y. enterocolitica O:9) has been tested with field bovine sera and found to improve the specificity of the rose bengal and complement fixation up to 70% (Weynants et al., 1996b; Weynants et al., 1996a). Immunological crossreactivities with proteins of taxonomically unrelated bacteria seem limited to some highly conserved proteins, such as heat shock protein-65 (Díaz and Bosseray, 1974; Spencer et al., 1994). On the other hand, they

428

E. Moreno and I. Moriyó n

are extensive with the proteins of taxonomically related bacteria, but although they can be shown with sera from naturally infected hosts (Drancourt et al., 1997; Velasco et al., 1997), they do not seem to represent a relevant source of nonspecificity (Velasco et al., 1997) and can be easily discriminated by using smooth LPS tests. Thus, Brucella protein antigens are perhaps the most useful antigens for the differential diagnosis of brucellosis and yersiniosis. Although they have been used in some antibody (Baldi et al., 1996) and lymphocyte blastogenesis tests (Chukwu, 1987), the skin test with proteins saline extracted from rough B. melitensis 115 (brucellin or brucellergene; Bhongbhibhat et al., 1970; Jones et al., 1973) is very useful and relatively simple to implement (Chukwu, 1987; Saegerman et al., 1999). There are contradictory reports on the value of the γ-interferon in the differential diagnosis of brucellosis and yersiniosis in cattle (Kittelberger et al., 1997; Weynants et al., 1995).

Treatment The ability of Brucella organisms to replicate within cell vacuoles is a determining factor in the chronic course of the disease and in the tendency to relapse. Within the intracellular niche, Brucella organisms are protected from a large number of antibiotics that cannot reach the intracellular milieu. Therefore, an appropriate therapy must use antibiotics capable of killing both extracellular and intracellular Brucella organisms (Ariza, 1999). Unfortunately, not a single antibiotic therapy achieves complete eradication of intracellular infection. In this respect, synergistic or additive effect by the use of two or more antibiotics for prolonged periods is required to cure and to reduce the frequency of relapses (Fig. 40). The possibility of enhancing the effectiveness of antibiotics showing good in vitro activity by including them into liposomes has been studied by several authors (Dees and Schultz, 1990; Vitas et al., 1997), but attempts to use some of these formulations in animals have been discouraging (Nicoletti et al., 1989). The development of antibiotic resistance by Brucella has never been demonstrated. For instance, the susceptibility of strains from human patients with relapses remains identical to that of strains isolated during the initial episode (Ariza et al., 1986); similarly, Brucella strains isolated from animals that did not clear the infection after the first antibiotic therapy show the same antibiotic susceptibility as that of the first isolates (Guerra and Nicoletti, 1986). The absence of plasmids (Genome Evolution) should be an important factor explaining, at least in part, the observation that the in vitro susceptibility of Brucella strains

CHAPTER 3.1.16 Antibiotics and days of treatment Doxycycline-streptomycin 45 Doxycycline-rifampicin 45 Tetracycline-streptomycin 30 Doxycycline-gentamycin 45 Doxycycline-netilmycin 45 Tetracycline-streptomycin 21 Tetracycline 42 Tetracycline 21 Doxycycline-rifampicin 30 Co-trimoxazole 45 0

10

20

30

40

Percentage of relapses Fig. 40. Relapse rate after antibiotic therapy of human brucellosis. Patients, naturally infected with Brucella organisms, were treated with a combination of antibiotics for different periods (indicated on the vertical axis). The period between the end of treatment and relapse and the percentage of relapses were registered, accordingly. Bars indicate average values. Standard error was less than 10% in all cases, except in doxycycline plus rifampicin treatment for 45 days in which standard error was close to 30%. Dr. Javier Ariza, from the Infectious Disease Service, Hospital De Bellvitge, Barcelona, Spain, supplied this information.

to antibiotics remains stable over time (Bosch et al., 1986). Human Brucellosis The antibiotic treatment of human brucellosis has been recently reviewed (Ariza, 1999), and only the most significant facts are summarized here. Since there is no antibiotic therapy effective in the totality of human cases, a careful follow-up of the patients is important. It is an established fact that tetracyclines are the most effective antibiotics in the treatment of brucellosis but also that by themselves they are not satisfactory and result in high rates (from 15–30%, depending on the duration of the treatment) of relapses. Monotherapy with other antibiotics, such as aminoglycosides, rifampicin, azithromycin, co-trimoxazol, or second generation fluoroquinolones has produced unsatisfactory results in laboratory models, human patients or both. Presently, a prolonged tetracycline-aminoglycoside combination (doxycycline 100 mg twice/day for 45 days and streptomycin 1 g/day [or 750 mg in patients older than 50 years] for 14 days) is the best treatment of the disease as judged by the comparatively low (3–5%) rates of relapse. Although aminoglycosides have low intracellular penetration, their synergy with tetracyclines is well established,

CHAPTER 3.1.16

and in most cases, their effectiveness in this combination offsets the problems (parenteral administration and ototoxicity). Some aminoglycosides such as gentamycin have better in vitro activity against Brucella and show lower toxicity, and they are currently recommended as streptomycin substitutes. Because of its better tolerance, doxycycline is the tetracycline used. It has to be emphasized that the duration of the treatment with both antibiotics is essential to obtain the best results. As an alternative to aminoglycosides, rifampicin can be used (900 mg in a single morning dose) along with doxycycline, both administered during 45 days. Although this treatment has no side effects and is better tolerated by the patient, its effectiveness is less than that of the doxycyclineaminoglycoside combination (the rate of relapses can be as high as 15%). It is, however, the therapy of choice when infections by B. melitensis Rev. 1 (Vaccines and Vaccination) occur or are suspected (Blasco and Díaz, 1993a). This sort of infection occurs most often in veterinarians and farmers, and there are also reports of accidental inhalation in vaccine-producing plants. In these cases, it is recommended that bacterial cultures be performed and the isolate typified or at least tested for streptomycin resistance. Brucellosis in pregnant women and in children less than seven years old requires a different therapy because of the side effects of the antibiotics of the standard treatment. There is no established optimal therapy, but rifampicin for 8 weeks in the first case and rifampicin for 4–6 weeks plus gentamycin for the first 7–10 days in children are recommended. Recent reports suggest that gentamycin is less effective than streptomycin in children (Issa and Jamal, 2000). Brucella endocarditis is a dangerous form of the disease that also calls for a special treatment to ensure clearance of the infection. Doxycycline plus gentamycin, both for 3 weeks, plus rifampicin for at least 8 weeks is recommended. Accidental inoculation or exposure to virulent Brucella in the diagnostic and research laboratories calls for a prophylactic antibiotic therapy. For this purpose, doxycycline alone is effective provided the treatment starts immediately. Although the duration of this treatment has not been determined, it is generally considered that the standard dose (doxycycline 100 mg twice/ day) for 7–10 days is effective in preventing the development of the disease. Animal Brucellosis Antibiotics have been rarely employed in animal brucellosis but they have been applied in some cases to treat animals of high genetic value or to prevent spreading of

The Genus Brucella

429

the disease in small dairy herds of highly valuable animals. In cattle, monotherapy with either oxytetracycline or free or liposomal streptomycin (Nicoletti et al., 1985; Nicoletti et al., 1989) is very unsatisfactory. The combination of long-acting oxytetracycline (for example, 20 mg/kg administered intramuscularly every 3 days for 21 days) and streptomycin or dihydostreptomycin (for example 20 mg/kg administered intramuscularly daily for 10 days) is successful in eliminating milk shedding of B. abortus in 55–71% of treated animals (Jiménez de Bagü és et al., 1991; Milward et al., 1984; Nicoletti et al., 1985; Nicoletti et al., 1989). A more prolonged treatment (14 oxytetracycline and 8 streptomycin doses) accompanied by intramammary infusion with oxytetracycline for 4 days is reported to have a 100% rate of success (Radwan et al., 1993). In the noncured animals, the oxytetracycline and streptomycin therapy reduces the number of bacteria in milk or in tissues after necropsy (Jiménez de Bagü és et al., 1991; Milward et al., 1984; Nicoletti et al., 1985; Nicoletti et al., 1989) and when combined with adult vaccination with B. abortus 19, allows for control of the disease without eliminating the infected animals (Jiménez de Bagü és et al., 1991). Antibiotic therapy also has been used in infections caused by B. ovis in rams. Like in human or cattle brucellosis, streptomycin or tetracyclines by themselves are not successful (Kuppuswamy, 1954; Marín et al., 1989a). On the other hand, combinations of aureomycin plus streptomycin (Kuppuswamy, 1954) or tetracyclines plus streptomycin have produced satisfactory results (Giauffret and Sanchis, 1974; Marín et al., 1989a). However, successful antibiotic therapy does not result in a clearance of the lesions characteristic of this disease (Marín et al., 1989a), and semen quality after treatment is reported to be poor (Kuppuswamy, 1954). Brucellosis in horses is quite severe and prolonged treatments are required. A mixture of oxytetracycline (20 mg/kg) and streptomycin (15 mg/kg) for at least 21 days is recommended.

Applications Similarly to other intracellular parasites such as Mycobacterium species, Brucella organisms and their products have been used as adjuvants, antitumoral agents, antiviral components and immunological tools as well as vectors for delivering foreign antigens. In addition, most of the Brucella species are primary pathogens capable of infecting healthy hosts and in consequence candidates to be used as biological weapons.

430

E. Moreno and I. Moriyó n

Brucella Vaccines as Vectors for Delivering Foreign Antigens Live bacterial vaccines are scarce and rather exceptional among useful vaccines. Mycobacterium BCG and attenuated Brucella vaccines developed more than 70 years ago are among the accepted vaccines currently in use. Brucella vaccines, such as B. abortus strain 19, B. melitensis Rev. 1 and B. abortus RB51, are the best known vaccines used in many countries for the control and eradication of bovine, caprine and ovine brucellosis. In contrast to the attenuated mycobacterial vaccine, the efficacy of which in open populations is controversial, Brucella vaccines have been efficiently used for many years (Vaccines and Vaccination). This recognized status makes Brucella vaccines good vector candidates for the expression and delivery of foreign antigens (Comerci et al., 1998; Vemulapalli et al., 2000). Brucella abortus strain 19 recombinant vaccine expressing a reporter repetitive protein of Trypanosoma cruzi induces good antibody responses against the alien protein in mice. In vitro and in vivo expression of this repetitive Trypanosoma antigen does not alter the Brucella growth pattern, nor does it generate a toxic lethal effect in the bacterium. Similarly, rough B. abortus RB51 vaccine has been used to express Mycobacterium heat shock protein 65. The foreign mycobacterial protein expressed in this vector vaccine induces a typical Th1-type immune response in mice, as indicated by the presence of IgG2 and INFγ, without the production of IgG1 or the secretion of IL-4. The immunogenic properties and growth conditions of this recombinant vector B. abortus RB51 remain without change as compared to the parental strain. The rationale for using Brucella vaccines as vectors for delivering foreign antigens is based on the assumption that these attenuated intracellular bacteria could successfully deliver antigens inside antigen-presenting cells (Splitter and Everlith, 1986; Splitter and Everlith, 1989). Since attenuated Brucella organisms follow the endocytic pathway (Escaping from the Endocytic Pathway), the prediction is that antigens delivered by this manner will follow the appropriate antigen processing route necessary for eliciting an efficient and protective immune response against foreign antigens. An important prediction would be that the expression of foreign antigens does not exclude the immune response to Brucella. In this respect, a double purpose vaccine would be developed.

Antitumoral and Antiviral Activity of Brucella Cells and Their Fractions Live and killed bacteria and subcellular Brucella fractions have been used experimentally as anti-

CHAPTER 3.1.16

tumoral reagents in mice and humans (Chirigos et al., 1978; Dazord et al., 1980; Dazord et al., 1984; Schultz et al., 1978). Trials have demonstrated that live Brucella vaccines, killed bacterins or insoluble cellular extracts (mainly containing peptidoglycan, outer-membrane fractions and ribosomes) are capable of inducing the reduction of tumoral masses without causing unfavorable reactions. Although insoluble Brucella extracts exert antitumoral activity, better results are achieved when irradiated tumor cells and Brucella extracts are administered at the same time. The antitumoral activity mediated by Brucella vaccines can be understood within the context of the Mackaness effect (T Cell Responses), in which transitional acquired cellular resistance against invaders (e.g., cancer cells) could be nonspecifically induced by Brucella organisms or other intracellular parasites, such as Mycobacterium. The logic behind the use of Brucella-human immunodeficiency virus (HIV) protein conjugates as vaccines is based on the property that Brucella bacterins have to behave as T-independent antigens (Antibody Response), which is a strategy to bypass the requirement for CD4+ cells in the antibody response of AIDS patients (Golding et al., 1995; Lapham et al., 1996). Killed Brucella bacterins conjugated with gp120 or V3 loop peptide derived from HIV induce neutralizing antibodies against HIV. BrucellaHIV protein conjugates mainly generate IgG2a primary and secondary responses in mice, even after CD4+ depletion. This type of response also has been observed with other foreign “academic” antigens conjugated to Brucella, such as trinitrophenyl, indicating that it is a general behavior of Brucella bacterins (Sacks and Kilnman, 1997). Sera from mice immunized with Brucella-gp120 display more potent anti-HIV activity than sera from mice immunized with the Brucella-V3 loop peptide. In addition, killed B. abortus stimulates Th1-type immunity with the production of INFγ and generation of virusspecific cytotoxic T-lymphocyte response in both normal and CD4+-depleted mice. In addition to the T-independent properties of Brucella bacterin (Antibody Response), the antibody and T cell cytotoxic responses are linked to the adjuvant effect displayed by Brucella cells (Brucella Bacterins and Envelope Molecules as Adjuvants). Brucella also promotes the secretion of other cytokines, such as IL-12, a molecule that acts synergistically with IL-2 to induce Th1 activation and differentiation of cytotoxic cells (Huang et al., 1999). Of course, the low endotoxic activity displayed by Brucella LPS (Moreno et al., 1981; Rasool, 1992) is a factor that may favor the use of Brucella bacterin-conjugates as adjuvants.

CHAPTER 3.1.16

Brucella Bacterins and Envelope Molecules as Adjuvants In contrast to enterobacterial LPS, Brucella LPS behaves as a T-independent type 1 carrier that stimulates the production of high levels of IgG2 and moderate levels of IgM in the endotoxinresistant C3H/HeJ mice strain (Moreno and Berman, 1979; Moreno et al., 1984b; Kurtz and Berman, 1986). The primary and even the stronger secondary responses induced by the Brucella LPS adjuvant activity could occur without the assistance of T cells. By taking advantage of these properties, together with the low endotoxicity displayed by Brucella LPS (Moreno et al., 1981; Rasool, 1992), this molecule has been used as an adjuvant in foot-and-mouth disease, for enhancing the immune response against the virus (Berinstein et al., 1993; Betts et al., 1993), and to potentiate antibody production against snake venom toxins (Rucavado et al., 1996) and has been proposed as an adjuvant for HIV vaccines (Goldstein et al., 1992). Murein-rich insoluble fractions have been experimentally used as adjuvant for generating antibodies against a variety of antigens, with variable success (Galdiero, 1995; Serre, 1982). It has been observed that these fractions induce quantitative and qualitative changes in the T and B lymphocyte population in mice, mainly with an initial increase of CD4+ and B cells, followed by an increase of CD8+ lymphocytes and a concomitant decrease of CD4+ and B cells.

Brucella Bacterins and Lipopolysaccharide as Immunological Tools Brucella cells bind to human B cells and to certain leukemic cells of B origin, forming rosettes (Lee et al., 1983; Teodorescu et al., 1977). This quality has been used for diagnostic purposes in the differentiation of leukemic cells. The Brucella B cell receptor has been associated with the LPS, since the M serotype is more prone than the A serotype to bind to B lymphocytes (Fig. 15; Opolysaccharide). Brucella LPS and trinitrophenylated-Brucella bacterin have been extensively used as T-independent antigens in immunological studies (Betts et al., 1993; Moreno et al., 1984b; Kurtz et al., 1986; Rennick et al., 1983; Tenay and Strober, 1985). Moreover, trinitrophenylated-Brucella bacterin has been regarded as one of the classical T-independent type-1 antigens and used as an academic model for demonstrating this type of antibody response dependent on B lymphocytes.

Brucella as a Biological Weapon Brucella species, particularly B. melitensis and B. suis, are potential agents of biological warfare.

The Genus Brucella

431

There are several properties that make these bacteria good candidates as biological weapons (Anonymous, 2000; Kaufmann et al., 1997). The main one is that members of the genus are primary pathogens inducing a debilitating and incapacitating severe disease that requires expensive and long-term treatment. It has been estimated that less than five bacteria of B. melitensis 16M are necessary to infect one human being. Alertness to suspected cases of biological terrorism with B. melitensis and investigations leading to integrated clinical, public health, and law enforcement against biological terrorism with Brucella strains have taken place in the United States. A model that compares the impact of Brucella melitensis released as aerosols in suburbs of a major city has been developed in this country. The model demonstrated that the economic impact of a bioterrorist attack in a populated area with B. melitensis could be close to $500 million per 100,000 persons. The model takes into account preventive measures, including rapid implementation of a post-attack prophylaxis program.

Literature Cited Abraham, W. R., C. Strompl, H. Meyer, S. Lindholst, E. R. Moore, R. Christ, M. Vancanneyt, B. J. Tindall, A. Bennasar, J. Smit, M. Tesar. 1999. Phylogeny and polyphasic taxonomy of Caulobacter species. Proposal of Maricaulis gen. nov. with Maricaulis maris (Poindexter) comb. nov. as the type species, and emended description of the genera Brevundimonas and Caulobacter. Int. J. Syst. Bacteriol. 49:1053–1073. Ackermann, H. W., F. Simon, and J. M. Verger. 1981. A survey of Brucella phages and morphology of new isolates. Intervirology 16:1–7. Ackermann, M. R., N. F. Cheville, and B. L. Deyoe. 1988. Bovine ileal dome lymphoepithelial cells: Endocytosis and transport of Brucella abortus strain 19. Vet. Pathol. 25:38–35. Adams, L. G. 1990. Development of live Brucella vaccines. In: L. G. Adams (Ed.) Advances in Brucellosis Research. Texas A&M University Press. College Station, TX. 250–276. Akira, S., H. Isshiki, T. Sugita, O. Tanabe, S. Kinoshita, Y. Nishio, T. Nakajima, T. Hirano, and T. Kishimoto. 1990. A nuclear factor for IL-6 expression (NF-IL6) is a member of a C/EBP family. EMBO J. 9:1897–1906. Alcantara, B., J. Gee, and R. M. Roop, II. 2000. Characterization of Brucella abortus mutants defective in survival under conditions of starvation and stress. In: D. O’Callaghan (Ed.) Brucellosis 2000. INSERM, Faculté de Médicine. Nîmes, France. 100:94. Alford, C. E., T. E. King Jr., and P. A. Campell. 1991. Role of transferrin, transferrin receptors, and iron in macrophage listericidal activity. J. Exp. Med. 174:459–466. Allardet-Servent, A., G. Bourg, M. Ramuz, M. Pages M., Bellis, and G. Roizes. 1988. DNA polymorphism in strains of the genus Brucella. J. Bacteriol. 170:4603– 4607.

432

E. Moreno and I. Moriyó n

Allen, C. A., L. G. Adams, and T. A. Ficht. 1998. Transposonderived Brucella abortus rough mutants are attenuated and exhibit reduced intracellular survival. Infect. Immun. 66:1008–1016. Almiró n, M., M. Matínez, N-Sanjuan; R. Ugalde. 2001. Ferrochelatase is present in Brucella abortus and is critical for its intracellular survival and virulence. Infect. Immun. 69 :6225–6230. Alnor, D., N. Frimodt-Mø ller, F. Espersen, and W. Frederiksen. 1994. Infections with unusual human pathogens Agrobacterium and Ochrobactrum anthropi. Clin. Infect. Dis. 18:914–920. Alonso-Urmeneta, B., I. Moriyó n, R. Díaz, and J. M. Blasco. 1988. Enzyme-linked immunosorbent assay with Brucella native hapten polysaccharide and smooth lipopolysaccharide. J. Clin. Microbiol. 26:2642–2646. Alpuche-Aranda, C. M., J. A. Swanson, W. P. Loomis, and S. I. Miller. 1992. Salmonella typhimurium activates virulence gene transcription within acidified macrophage phagosomes. Proc. Natl. Acad. Sci. USA 89:10079– 10083. Altenbern, R. A., and R. D. Housewright. 1951. Alanine synthesis and carbohydrate oxidation by smooth Brucella abortus. J. Bacteriol. 62:97–105. Altenbern, R. A., and R. D. Housewright. 1953. Carbohydrate oxidation and citric synthesis by smooth Brucella abortus 19. Arch. Biochem. Biophys. 36:345–356. Alton, G. G., and S. S. Elberg. 1967. Rev. 1 Brucella melitensis vaccine: A review of ten years of study. Vet. Bull. 371:793–800. Alton, G. 1977. Experience with Brucella vaccines. In: R. P. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 209– 217. Alton, G. G., and S. S. Elberg. 1981. Rev 1 Brucella melitensis vaccine. Part II: 1968–1980. Vet. Bull. 51:67–73. Alton, G. G., L. M. Jones., R. D. Agnus, and J. M. Verger. 1988. Techniques for the Brucellosis Laboratory. Institut National de la Recherche Agronomique. Paris, France. Alton, G. G. 1990a. Brucella melitensis. In: K. H. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 383–409. Alton, G. G. 1990b. Brucella suis. In: K. H. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 411–422. Alvárez-Domínguez, C., and P. Stahl. 1999. Increased expression of rab5a correlates directly with accelerated maturation of Listeria monocytogenes phagosomes. J. Biol. Chem. 274:11459–11462. Amábile-Cuevas, C. F., and M. E. Chicurel. 1992. Bacterial plasmids and gene flux. Cell 70:189–199. Amano, K., M. Fujita, T. Suto. 1993. Chemical properties of lipopolysaccharides from spotted fever group rickettsiae and their common antigenicity with lipopolysaccharides from Proteus species. Infect. Immun. 61:4350–4355. Amano, K. I., J. C. Williams, G. A. Dasch. 1998. Structural properties of lipopolysaccharides from Rickettsia typhi and Rickettsia prowazekii and their chemical similarity to the lipopolysaccharide from Proteus vulgaris OX19 used in the Weil-Felix test. Infect. Immun. 66:923–926. Anacker, R. L., R. A. Finkelstein, W. J. Haskins, M. Landy, K. L. Milner, E. Ribi, and P. W. Sashk. 1964. Origin and properties of naturally ocurring hapten from Escherichia coli. J. Bacteriol. 88:1705–1720. Anast, N.; J. Smit. 1988. Isolation and characterization of marine caulobacters and assessment of their potential

CHAPTER 3.1.16 for genetic experimentation A. Environ. Microbiol. 54:809–817. Andersen, P. S., P. M. Lavoie, R. P. Sekaly, H. Churchill, D. M. Kranz, P. M. Schlievert, K. Karjalainen, and R. A. Mariuzza. 1999. Role of the T cell receptor alpha chain in stabilizing TCR-superantigen-MHC class II complexes. Immunity 10:473–483. Anderson, J. D., and H. Smith. 1965. The metabolism of erythrithol by Brucella abortus. J. Gen. Microbiol. 38:109–124. Anderson, T. D., and N. F. Cheville. 1986a. Ultrastructural morphometric analysis of Brucella abortus-infected trophoblasts in experimental placentitis. Bacterial replication occurs in rough endoplasmic reticulum. Am. J. Pathol. 124:226–237. Anderson T. D., N. F. Cheville, and V. P. Meador. 1986b. Pathogenesis of placentitis in the goat inoculated with Brucella abortus. II: Ultrastructural studies. Vet. Pathol. 23:227–239. Anderson, T. D., V. P. Meador, and N. F. Cheville. 1986c. Pathogenesis of Placentitis in the goat inoculated with Brucella abortus. I: Gross and histologic lesions. Vet. Pathol. 23:219–226. Andersson, J. O., and S. G. Andersson. 1997. Genomic rearengments during evolution of obligate intracellular parasite Rickettsia prowasekii as inferred from an analysis of 52015 bp nucleotide sequence. Microbiology 143:2783–2795. Andersson S. G., A. Zomorodipour, J. O. Andersson, T. Sicheritz-Ponten, U. C. Alsmark, R. M. Podowski, A. K. Naslund, A. S. Eriksson, H. H. Winkler, C. G. Kurland. 1998. The genome sequence of Rickettsia prowazekii and the origin of mitochondria. Nature 396:133–140. Andersson, S. G., C. Dehio. 2000. Rickettsia prowasekii and bartonella henselae: differences in the intracellular life styles revised. Int. J. Med. Microbiol. 290 :135–141. Andrejewski, N., E. L. Punnonen, G. Guhde, Y. Tanaka, R. Lullmann-Rauch, D. Hartmann, K. Von-Figura, and P. Saftig. 1999. Normal lysosomal morphology and function in LAMP-1-deficient mice. J. Biol. Chem. 274:12692–12701. Anonymous. 2000. Suspected brucellosis case prompts investigation of possible bioterrorism-related activity— New Hampshire and Massachusetts, 1999. Morbidity & Mortality Weekly Report 49:509–12. Antoine, J., E. Prina, C. Jouane, and P. Bongrand. 1990. Parasitophorous vacuoles of Leishmania amazonensisinfected macrophages maintain an acidic pH. Infect. Immun. 58:779–787. Aragó n, V. 1996a. Caracterizació n bioquímica y propiedades inmunoló gicas de los polisacáridos haptenicos de Brucella (PhD thesis). University of Navarra. Navarra, Spain. 1–163. n. 1996b. Aragó n, V., R. Díaz, E. Moreno, and I. Moriyó Characterization of Brucella abortus and Brucella melitensis native haptens as outer-membrane O-type polysaccharides independent from the smooth lipopolysaccharide. J. Bacteriol. 178:1070–1079. Araya, L. N., P. H. Elzer, G. E. Rowe, F. M. Enright, and A. J. Winter. 1989. Temporal development of protective cell-mediated and humoral immunity in BALB/c mice infected with Brucella abortus. J. Immunol. 143:3330– 3337. Araya, L. N., and A. J. Winter. 1990. Comparative protection of mice against virulent and attenuated strains of Bru-

CHAPTER 3.1.16 cella abortus by passive transfer of immune T cells or serum. Infect. Immun. 58:254–256. Arenas, G. N., A. S. Staskevich, A. Aballay, and L. S. Mayorga. 2000. Intracellular trafficking of Brucella abortus in mice. Infect. Immun. 68:4255–4263. Arese, A., S. Cravero, M. Boschiroli, E. Campos, L. Samartino, and O. L. Rossetti. 1999. Uso de una proteína recombinante de Brucella abortus para el diagnostico de la brucelosis en diferentes especies animales. Rev. Argent. Microbiol 31 (Suppl. 1):36–39. Ariza, J., J. Bosch, F. Guidiol, J. Liñ ares, P. FernándezViladrich, and R. Martín. 1986. Relevance of in vitro antimicrobial susceptibility of Brucella melitensis to relapse rate in human brucellosis. Antimicrob. Agents Chemother. 30:958–960. Ariza, J., T. Pellicer, R. Pallares, A. Foz, and F. Gudiol. 1992. Specific antibody profile in human brucellosis. Clin. Infect. Dis. 14:131–140. Ariza, J. 1998. Brucelosis: Perspectiva actual de la enfermedad. Perdil de las immunoglobulinas en el curso de la evolució n (PhD thesis). Facultad de Medicina, Universidad Autó noma de Barcelona. Barcelona , Spain. 1– 87. Ariza, J. 1999. Brucellosis: An update. The perspective from the mediterranean basin. Rev. Med. Microbiol. 10:125– 136. Asarta, A. 1989. Erradicació n de la brucelosis en el ganado vacuno de Navarra. In: Sociedad Españ ola de Microbiología Actas del XII Congreso Nacional de Microbiología. SEM. Pamplona, Spain. 371–373. Avise, J. C., and R. M. Ball. 1990. Principles of genealogical concordance in species concepts and biological taxonomy. Oxford Surv. Evol. Biol. 7:45–67. Awram, P., and J. Smit. 2001. Identification of lipopolysaccharide O antigen synthesis genes required for attachment of the S-layer of Caulobacter crescentus. Microbiology 147:1451–1460. Baily, G. G., J. B. Krahn, B. S. Drasar, and N. G. Stoker. 1992. Detection of Brucella melitensis and Brucella abortus by DNA amplification. J.Trop. Med. Hyg. 95:271–275. Baldi, P. C., G. H. Giambartolomei, F. A. Goldbaum, L. F. Abdon, C. A. Velikovsky, R. Kittelberger, and C. A. Fossati. 1996. Humoral immune response against lipopolysaccharide and cytoplasmic proteins of Brucella abortus in cattle vaccinated with B. abortus S19 or experimentally infected with Yersinia enterocolitica serotype 0:9. Clin. Diagn. Lab. Immunol. 3:472–476. Baldwin, C. L., D. F. Antczak, and A. J. Winter. 1984. Evaluation of lymphocyte blastogenesis for diagnosis of bovine brucellosis. Dev. Biol. Stand. 56:357–369. Baldwin, C. L., and A. J. Winter. 1994. Macrophages and Brucella. Immunol. Ser. 60:363–380. Barker, J., P. A. Lambert, and M. R. Brown. 1993. Influence of intra-amoebic and other growth conditions on the surface properties of Legionella pneumophila. Infect. Immun. 61:3503–3510. Barnes, P. F., C. L. Grisso, J. S. Abrams, H. Band, T. H. Rea, and R. L. Modlin. 1992. γδ T lymphocytes in human tuberculosis. J. Infect. Dis. 5:506–510. Batrakov, S. G., D. I. Nikitin, V. I. Sheichenko, A. O. Ruzhitsky. 1997. Unusual lipid composition of the gram-negative, freshwater, stalked bacterium Caulobacter bacteroides NP-105. Biochim. Biophys. Acta. 1347:127–139. Beck, B. L., L. B. Tabatabai, and E. Mayfield. 1990. A protein isolated from Brucella abortus is a Cu-Zn superoxide dismutase. Amer. Chem. Soc. 29:372–376.

The Genus Brucella

433

Bellaire, B. H., P. H. Elzer, C. L. Baldwin, and R. M. Roop. 1998. Contribution of the Brucella abortus 2,3-dihydroxybenzoic acid (DHBA) biosynthesis operon to virulence in ruminants. Abstract 40 In: Proceedings 79th Annual Conference of Research Workers on Animal Diseases. 43. Bellaire, B. H., P. H. Elzer, C. L. Baldwin, R. M. Roop. 1999. The siderophore 2,3-dihydroxybenzoic acid is not required for virulence of Brucella abortus in BALB/c mice. Infect. Immun. 67:2615–2618. Bentejac, M. C., G. Biron, A. Bertrand, and S. Bascoul. 1984. Vaccination contre la brucellose humaine: Bilan sur une periode de 2 ans. Dev. Biol. Stand. 56:531–535. Bercovich, Z., and R. Taaijke. 1990. Enzyme immunoassay using mouse monoclonal anti-bovine antibodies for the detection of Brucella abortus antibodies in cow milk. Zentralbl. Veterinarmed. B 37:753–759. Berinstein, A., M. Perez-Filgueira, A. Schudel, P. Zamorano, M. Borca, and A. Sadir. 1993. Avridine and LPS from Brucella ovis: Effect on the memory induced by footand-mouth disease virus vaccination in mice. Vaccine 11:1295–1301. Bertotto, A., R. Gerli, F. Spinozzi, C. Muscat, F. Scalise, G. Castellucci, M. Sposito. F. Candio, and R. Vaccaro. 1993. Lymphocytes bearing the gd T cell receptor in acute Brucella melitensis infection. Eur. J. Immunol. 23:1177–1182. Bertram, T. A., P. C. Canning, and J. A. Roth. 1986. Preferential inhibition of primary granule release from bovine neutrophils by a Brucella abortus extract. Infect. Immun. 52:285–292. Betts, M., P. Beining, M. Brunswick, J. Inman, R. D. Angus, T. Hoffman, and B. Golding. 1993. Lipopolysaccahride from Brucella abortus behaves as a T-cell-independent type carrier in murine antigen-specific antibody responses. Infect. Immun. 61:1722–1729. Bhat, U. R., R. W. Carlson, M. Busch, and H. Mayer. 1991. Distribution and phylogenetic significance of 27hydroxy-octacosanoic acid in lipopolysaccharides from bacteria belonging to the alpha-2 subgroup of Proteobacteria. Int. J. Syst. Bacteriol. 41:213–217. Bhongbhibhat, N., S. S. Elberg, and T. H. Chen. 1970. Characterization of brucella skin-test antigens. J. Infect. Dis. 122:70–82. Birtles, R. J., T. G. Harrison, N. A. Saunders, and D. H. Molyneux. 1995. Proposal to unify the genera Grahamella and Bartonella with description of Bartonella talpae comb. nov., Bartonella peromysci comb. nov., and the new species Bartonella grahamii sp. nov., Bartonella taylorii sp. nov., and Bartonella doshiae sp. nov. Int. J. Syst. Bacteriol. 45:1–8. Blasco, J. M., R. Díaz, I. Moriyó n, and M. D. Salvo. 1984a. Evaluation of a radial immunodiffusion test for diagnosing brucellosis in sheep and its possible value for differentiating infected from Brucella melitensis REV 1 vaccinated sheep. Dev. Biol. Stand. 56:507–511. Blasco, J. M., A. Estrada, and M. Mercadal. 1984b. A note on adult sheep vaccination with reduce dose of Brucella melitensis. Ann. Rech. Vet. 15:554–556. n, and R. Blasco, J. M., M. Marín. M. Barberán. I. Moriyó Díaz. 1987. Immunization with Brucella melitensis Rev1 against B. ovis infection in ram. Vet. Microbiol. 14:381– 392. Blasco, J. M. 1990. B. ovis. In: K. Nielsen and B. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 351–378.

434

E. Moreno and I. Moriyó n

Blasco, J. M., and R. Díaz. 1993a. Brucella melitensis Rev-1 vaccine as a cause of human brucellosis. Lancet 342:805. Blasco, J. M., C. M. Marín, M. P. Jiménez de Bagü és, and M. Braberán. 1993b. Efficacy of Brucella suis strain 2 vaccine against Brucella ovis in rams. Vaccine 11:1291– 1294. Blasco, J. M., B. Garin-Bastuji, C. M. Marín, G. Gerbier, J. Fanlo, M. P. Jiménez de Bagü és, and C. Cau. 1994a. Efficacy of different rose bengal and complement fixation antigens for the diagnosis of Brucella melitensis infection in sheep and goats. Vet. Rec. 134:415–420. Blasco, J. M., C. M. Marín, M. P. Jiménez de Bagü és, M. Braberán, A. Hernandez, L. Molina, J. Velasco, R. Díaz, and I. Moriyó n. 1994b. Evaluation of allergic and serological tests for diagnosing Brucella melitensis infection in sheep. J. Clin. Microbiol. 32:1835–1840. Blasco, J. M. 1997. A review of the use of B. melitensis Rev 1 vaccine in adult sheep and goats. Prev. Vet. Med. 31:275–283. Bohin, J. P. 2000. Osmoregulated periplasmic glucans in Proteobacteria. FEMS Microbiol. Lett. 186:19. Bosch J., J. Liñ ares, M. J. Ló pez de Goicochea, J. Ariza, M. Cisnal, and R. Martín. 1986. In vitro activity of ciprofloxacin, ceftriaxone and five other antimicrobial agents against 95 strains of Brucella melitensis. J. Antimicrob. Chemother. 17:459–461. Boschiroli, M. L., S. Ouahrani-Bettache; V. Foulongne, S. Michaux-Charachon, G. Bourg, A. Allardet-Servent, C. Cazevieille, J. P. Liautard, M-Ramuz D. O’Callaghan. 2002. The Brucella suis virB operon is induced inttracellularly in macrophages. Proc. Nat. Acad. Sci. USA 99:1544–1549. Bosseray, N. 1983. Vaccine and serum-mediated protection against Brucella infection of mouse placenta. Br. J. Exp. Pathol. 64:617–618. Bosseray, N. 1984. Infection du plancenta de la souris par Brucella: Pathogénie es immunite. Dev. Biol. Stand. 56:283–287. Bosseray, N. 1991. Brucella melitensis Rev. 1 living attenuated vaccine: Stability of markers, residual virulence and immunogenicity in mice. Biologicals 19:355–363. Bouzar, H., D. Ouadah, Z. Krimi, J. B. Jones, M. Trovato, A. Petit, and Y. Dessaux. 1993. Correlative association between resident plasmids and the host chromosome in a diverse Agrobacterium soil population. Appl. Environ. Microbiol. 59:1310–1317. Bowden, R. A., A. Cloeckaert, M. Zygmunt, S. Bernard, and G. Dubray. 1995. Surface exposure of outer membrane protein and lipopolysaccharide epitopes in Brucella species studied by enzyme-linked immunosorbent assay and flow cytometry. Infect. Immun. 63:3945–3952. Boyd, C. 1977. Management and prevention. In: R. P. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 267– 331. Brade, H., L. Brade, and E. T. Rietschel. 1988. Structureactivity relationships of bacterial lipopolysaccharides (endotoxins). Zentralbl. Bakt. Hyg. A268:151–179. Breedveld, M. W., and K. J. Miller. 1994. Cyclic β-glucans of members of the family Rhizobiceae. Microbiol. Rev. 58:145–161. Brenner, D. J., D. G. Hollis, C. W. Moss, C. K. English, G. S. Hall, J. Vincent, J. Radosevic, K. A. Birkness, W. F. Bibb, F. D. Quinn, B. Swaminathan, R. E. Weaver, M. W. Reeves, S. P. O’Connor, P. S. Hayes, F. C. Tenover, A. G. Steigerwalt, B. A. Perkins, M. I. Daneshvar, B. C. Hill,

CHAPTER 3.1.16 J. A. Washington, T. C. Woods, S. B. Hunter, T. L. Hadfield, G. W. Ajello, A. F. Kaufmann, D. J. Wear, and J. D. Wenger. 1991. Proposal of Afipia gen. nov., with Afipia felis sp. nov. (formerly the cat scratch disease bacillus), Afipia clevelandensis sp. nov. (formerly the Cleveland Clinic Foundation strain), Afipia broomeae sp. nov., and three unnamed genospecies. J. Clin. Microbiol. 29:2450–2460. Brenner, D. J., S. P. O’Connor, H. H. Winkler, and A. G. Steigerwalt. 1993. Proposals to unify the genera Bartonella and Rochalimaea, with descriptions of Bartonella quintana comb. nov., Bartonella vinsonii comb. nov., Bartonella henselae comb. nov., and Bartonella elizabethae comb. nov., and to remove the family Bartonellaceae from the order Rickettsiales. Int. J. Syst. Bacteriol. 43:777–786. Bress, D., E. Steadham, M. Stevens, and N. Cheville. 1996. α5β1 integrin mediates Brucella abortus strain RB51 adhesion on bovine trophoblastic cells. Vet. Pathol. 33:615. Bricker, B. J., L. B. Tabatabai, B. A. Judge, B. L. Deyoe, and J. E. Mayfield. 1990. Cloning, expression, and occurrence of the Brucella Cu-Zn superoxide dismutase. Infect. Immun. 58:2935–2939. Bricker, B. J., and S. M. Halling. 1994. Differentiation of Brucella abortus bv. 1, 2, and 4, Brucella melitensis, Brucella ovis, and Brucella suis bv. 1 by PCR. J. Clin. Microbiol. 32:2660–2666. Bricker, B. J., and S. M. Halling. 1995. Enhancement of the Brucella AMOS PCR assay for differentiation of Brucella abortus vaccine strains S19 and RB51. J. Clin. Microbiol. 33:1640–1642. Bricker, B. J. 1999. Differentiation of hard-to-type bacterial strains by RNA mismatch cleavage. Biotechniques 27:321–326. Bricker, B. J., D. R. Ewalt, A. P. MacMillan, G. Foster, and S. Brew. 2000. Molecular characterization of Brucella strains isolated from marine mammals. J. Clin Microbiol. 38:1258–1262. Briones, G., N. Iñ on de Iannino, M. Steinberg, and R. Ugalde. 1997. Periplasmic cyclic 1,2-beta-glucan in Brucella spp. is not osmoregulated. Microbiology 143:1115–1124. Brodie, J., and G. P. Sinton. 1975. Fluid and solid media for isolation of Brucella abortus. J. Hyg. (Lond.) 74:359–367. Brown, G. M., C. R. Ranger, and D. J. Kelley. 1971. Selective media for the isolation of Brucella ovis. Cornell. Vet. 61:265–280. Brun, M., and S. Descous. 1984. Homogénéité électrophorétique de l’urease chez 3 espèces de Brucella. Dev. Biol. Stand. 56:113–121. Brunham, R. C., F. A. Plummer, and R. S. Stephens. 1993. Bacterial antigenic variation, host Immune response, and pathogen-host coevolution. Infect. Immun. 61:2273– 2276. Buchmeier, N. A., and F. Heffron. 1990. Induction of Salmonella stress proteins upon infection of macrophages. Science 248:730–732. Burgdorfer, W. 1967. Trans-stadial and transovarial development of disease agents in arthropods. Ann. Rev. Entomol. 12:347–376. Burns, B. P., S. L. Hazell, and G. L. Mendz. 1995. Acetyl-CoA carboxylase activity in Helicobacter pylori and the requirement of increased CO2 for growth. Microbiology 141:3113–3118. Cameron, H. S., B. Holmes, and M. E. Meyer. 1952. Comparative metabolic studies on the genus of Brucella. I. Evi-

CHAPTER 3.1.16 dence of a urea cycle from glutamic acid metabolism. J. Bacteriol. 64:709–712. Cameron, H. S., and M. E. Meyer. 1954. Comparative metabolic studies on the genus Brucella. II. Metabolism of amino acids that occur in the urea cycle. J. Bacteriol. 67:34–37. Campbell, G. A., L. G. Adams, and B. A. Sowa. 1994. Mechanism of binding of Brucella abortus to mononuclear phagocytes from cows naturally resistant or susceptible to brucellosis. Vet. Immunol. Immunopathol. 41:295– 306. Campbell, G. R., Reuhs, and G. C. Walker. 2002. Chronic intracellular infection of alfalfa nodules by Sinorhizobium meliloti requires correct lipopolysaccharide. Proc. Nat. Acad. Sci. USA 99 :3938–3943. Canback, B., S. G. E. Andersson, and C. G. Kurland. 2002. The global phylogeny of glycolytic enzymes. Proc. Nat. Acad. Sci. USA 99:6097–6102. Canning, P. C., J. A. Roth, and B. L. Deyoe. 1986. Release of 5′-guanosine monophosphate and adenine by Brucella abortus and their role in the intracellular survival of the bacteria. J. Infect. Dis. 154:464–470. Canning, P. C., B. L. Deyoe, and J. A. Roth. 1988. Opsonindependent stimulation of bovine neutrophil oxidative metabolism by Brucella abortus. Amer. J. Vet. Res. 49:160–163. Carmichael, L. E., S. J. Zoha, and R. Flores-Castro. 1984. Problems in the serodiagnosis of canine brucellosis: Dog responses to cell wall and internal antigens of Brucella canis. Dev. Biol. Stand. 56:371–376. Carmichael, L. E. 1990. Brucella canis. In: K. H. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 336–350. Carmichael, L. E., and S. J. Shin. 1996. Canine brucellosis: A diagnostician’s dilemma. Semin. Vet. Med. Surg. Small. Anim. 11:161–165. Caron, E. J., M. Cellier, P. Liautard, and S. Kohler. 1994a. Complementation of a DnaK-deficient Escherichia coli strain with dnaK/dnaJ operon of Brucella ovis reduces the rate of initial intracellular killing within the monocytic cell line U937. FEMS Microbiol. Lett. 120:335–340. Caron, E., J. P. Liautard, and S. Kohler. 1994b. Differentiated U937 cells exhibit increased bactericidal activity upon LPS activation and discriminate between virulent and avirulent Listeria and Brucella species. J. Leuk. Biol. 56:174–181. Caron, E., A. Gross, J.-P. Liautard, and J. Dornan. 1996. Brucella species release a specific, protease-sensitive inhibitor of TNF-α expression active on human macrophage-like cells. J. Immunol. 257:2885–1891. Casas, J., Y. Partal, J. Llosa, J. Leiva, J. M. Navarro, and R. M. de la Fuente. 1994. Detecció n de Brucella por un sistema automático de hemocultivos: Bact/Alert. Enferm. Infec. Microbiol. Clin. 12:497–500. Castañ eda, M. R. 1961. Laboratory diagnosis of brucellosis in man. Bull. WHO 24:73–84. Cella, M., A. Engering, V. Pinet, J. Pieters, and A. Lanzavecchia. 1997. Inflammatory stimuli induce accumulation of MHC class II complexes on dendritic cells. Nature 388:782–785. Cellier, M. F., J. Teyssier, M. Nicolas, J. P. Liautard, J. Marti, and W. J. Sri. 1992. Cloning and characterization of the Brucella ovis heat shock protein DnaK functionally expressed in Escherichia coli. J. Bacteriol. 174:8036– 8042.

The Genus Brucella

435

Cetinkaya, B., H. Ongor, A. Muz, H. B. Ertas, H. Kalender, and H. M. Erdogan. 1999. Detection of Brucella species DNA in the stomach content of aborted sheep fetuses by PCR. Vet. Rec. 144:239–240. Chaves-Olarte, E., C. Guzmán-Verri, Stephane Méresse, M. Desjardins, J. Pizarro-Cerda, J. Badilla, J-P. Gorvel, E. Moreno. 2002. Activation of Rho and Rab GTPases dissociate Brucella abortus internalization from intracellular trafficking. Cell. Microbiol. (in press). Cheers, C., H. Pavlov, C. Riglar, and E. Madraso. 1980. Macrophage activation during experimental murine brucellosis. III: Do macrophages exert feedback control during brucellosis? Cell. Immunol. 49:168–174. Cheers, C. 1984. Pathogenesis and cellular immunity in experimental murine brucellosis. Dev. Biol. Stand. 56:237–242. Cheng, H. P., and G. C. Walker. 1998. Succinoglycan production by Rhizobium meliloti is regulated through the ExoS-ChvI two component regulatory system. J. Bacteriol. 180:20–26. Cherwonogrodzky, J. W., and K. H. Nielsen. 1988. Brucella abortus 1119-3 O-chain polysaccharide to differentiate sera from B. abortus S-19-vaccinated and field-straininfected cattle by agar gel immunodiffusion. J. Clin. Microbiol. 26:1120–1123. Cherwonogrodzky, J., G. Dubray, E. Moreno, and H. Mayer. 1990. Antigens of Brucella. In: K. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 19–64. Chester, B., and L. H. Cooper. 1979. Achromobacter species (CDC group Vd): Morphological and biochemical characterization. J. Clin. Microbiol. 9:425–436. Cheville, N. F., A. E. Jensen, S. M. Halling, F. M. Tatum, D. C. Morfitt, S. G. Hennager, W. M. Frerichs, and G. Schurig. 1992. Bacterial survival, lymph node changes, and immunologic responses of cattle vaccinated with standard and mutant strains of Brucella abortus. Amer. J. Vet. Res. 53:1881–1888. Cheville, N. F., M. G. Stevens, A. E. Jensen, F. M. Tatum, and S. M. Halling. 1993. Immune responses and protection against infection and abortion in cattle experimentally vaccinated with mutant strains of Brucella abortus. Amer. J. Vet. Res. 54:1591–1597. Cheville, N. F., S. C. Olsen, A. E. Jensen, M. G. Stevens, A. M. Florance, H. H. Houng, E. S. Drazer, R. L. Warren, T. L. Hadfield, and D. L. Hoover. 1996a. Bacterial persistence and immunity of goats vaccinated with purE deletion mutant or the parental 16M strain of Brucella melitensis. Infect. Immun. 64:241– 2439. Cheville, N. F., S. C. Olsen, A. E. Jensen, M. G. Stevens, M. V. Palmer, and A. M. Florance. 1996b. Effects of age at vaccination on efficacy of Brucella abortus strain RB51 to protect cattle against brucellosis. Amer. J. Vet. Res. 57:1153–1156. Chirigos, M. A., W. A. Stylos. R. M. Schultz, and J. R. Fullen. 1978. Chemical and biological adjuvants capable of potentiating tumor cell vaccine. Cancer Res. 38:1085– 1091. Cho, H. J., and L. Niilo. 1987. Diagnostic sensitivity and specificity of an enzyme-linked immunosorbent assay for the diagnosis of Brucella ovis infection in rams. Can. J. Vet. Res. 51:99–103. Choma, A., T. Urbanik-Sypniewska, R. Russa, J. Kutkowska, and H. Mayer. 2000. Ocurrence and taxonomic significance of oxo-fatty acids in lipopolysaccharides from

436

E. Moreno and I. Moriyó n

members of Mesorhizobium. Syst. A. Microbiol. 23:185– 190. Choudhary, M., C. Mackenzie, K. S. Nereng, E. Sodergren, G. M. Weinstock, and S. Kaplan. 1995. Multiple chromosomes in bacteria: Structure and function of chromosome II of Rhodobacter sphaeroides 2.4.1(T). J. Bacteriol. 176:7694–7702. Christie, P. J., and A. Covacci. 2000. Bacterial type IV secretion systems; DNA conjugation machines adapted to export virulent factors. In: P. Cossart, P. Bouquet, S. Normark, and R. Rappuoli (Eds) Cellular Microbiology. American Society for Microbiology. Washington DC, 265–273. Chukwu, C. C. 1985. The instability of Brucella abortus strain 45/20 and a note on significance of using an unstable rough strain in the diagnosis of bovine brucellosis. Int. J. Zoonoses 12:120–125. Chukwu, C. C. 1987. Differentiation of Brucella abortus and Yersinia enterocolitica serotype 09 infections in cattle: The use of specific lymphocyte transformation and brucellin skin tests. Vet. Q. 9:134–142. Clavareau, C., V. Wellemans, K. Walravens, M. Tryland, J. M. Verger, M. Grayon, A. Cloeckaert, J. J. Letesson, and J. Godfroid. 1998. Phenotypic and molecular characterization of a Brucella strain isolated from a minke whale (Balaenoptera acutorostrata). Microbiology 144:3267– 3273. Cline, M. J., K. L. Melmon, W. C. Davies, and H. E. William. 1968. Mechanism of endotoxin interaction with human leukocytes. Br. J. Haematol. 15:539–547. Cloeckaert, A., P. de Wergifosse, G. Dubray, and J. N. Limet. 1990. Identification of seven surface-exposed Brucella outer membrane proteins by use of monoclonal antibodies: Immunogold labeling for electron microscopy and enzyme-linked immunosorbent assays. Infect. Immun. 58:3980–3987. Cloeckaert, A., P. Kerkhofs, and J. N. Limet. 1992. Antibody response to Brucella outer membrane proteins in bovine brucellosis: Immunoblot analysis and competitive enzyme-linked immunosorbent assay using monoclonal antibodies. J. Clin. Microbiol. 30:3168–3174. Cloeckaert, A., J. M. Verger, M. Grayon, and O. Grepinet. 1995. Restriction site polymorphism of the genes encoding the major 25 kDa and 36 kDa outer-membrane proteins of Brucella. Microbiology 141:2111–2121. Cloeckaert, A., J. M. Verger, M. Grayon, and O. Grepinet. 1996a. Polymorphism at the dnaK locus of Brucella species and identification of a Brucella melitensis speciesspecific marker. J. Med. Microbiol. 45:200–205. Cloeckaert, A., J. M. Verger, M. Grayon, and N. Vizcaíno. 1996b. Molecular and immunological characterization of the major outer membrane proteins of Brucella. FEMS Microbiol. Lett. 145:1–8. Cloeckaert, A., A. Tibor, and M. S. Zygmunt. 1999. Brucella outer membrane lipoproteins share antigenic determinants with bacteria of the family Rhizobiaceae. Clin. Diag. Lab. Immunol. Nouzilly, France 6:627–629. Cloeckaert, A., M. Grayon, J. M. Verger, J. J. Letesson, and F. Godfroid. 2000. Conservation of seven genes involved in the biosynthesis of the lipopolysaccharide O-side chain in Brucella spp. Res. Microbiol. 151:209–216. Comerci, D. J., G. D. Pollevick, A. M. Vigliocco, A. C. C. Frasch, and R. A. Ugalde. 1998. Vector Development for the Expression of Foreing Protein in the Vaccine Strain Brucella abortus S19. Infect. Immun. 66:3862– 3866.

CHAPTER 3.1.16 Comerci, D. J., M. J. Martínez-Lorenzo, R. Sieira, J.-P. Gorvel, and R. A. Ugalde. 2001. Essential role of the VirB machinery in the maturation of the Brucella abortuscontaining vacuole. Cell. Microbiol. 3:159–168. Corbeil, L. B., K. Blau, T. J. Inzana, K. H. Nielsen, R. H. Jacobson, R. R. Corbeil, and A. J. Winter. 1988. Killing of Brucella abortus by bovine serum. Infect. Immun. 56:3251–3261. Corbel, M. J., and E. L. Thomas. 1980a. The Brucella-phages: Their Properties, Characterization and Applications. Ministry of Agriculture, Fisheries and Food. London, UK. Booklet 2266:1–45. Corbel, M. J., A. C. Scott, and H. M. Ross. 1980b. Properties of a cell-wall-defective variant of Brucella abortus of bovine origin. J. Hyg. (Lond.) 85:103–113. Corbel, M. J., and W. J. Brinley-Morgan. 1984. Genus Brucella. In: N. R. Krieg and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 1:377–388. Corbel, M. J., and D. M. Hendry. 1985. Urease activity of Brucella species. Res. Vet. Sci. 38:252–253. Corbel, M. J. 1989. Microbiology of the genus Brucella. In: E. J. Young, and M. J. Corbel (Eds.) Brucellosis: Chemical and Laboratory. CRC Press. Boca Raton, FL. 53– 69. Correa, O. S. E. A. Rivas, and A. J. Barneix. 1999. Cellular envelopes and tolerance to acid pH in Mesorhizobium loti. Curr. Microbiol. 38:329–334. Cover, T. L., W. Puryear, G. I. Perez-Perez, and M. J. Blaser. 1991. Effect of urease on HeLa cell vacuolation induced by Helicobacter pylori cytotoxin. Infect. Immun. 59:1264–1270. Cracraft, J. 1983. Species concept and speciation analysis. In: R. F. Johnson (Ed.) Current Ornithology. Plenum Press. New York, NY. 159–187. Crawford R. P., and R. J. Hidalgo. 1977. Bovine Brucellosis. Texas A&M University Press. College Station , TX. 1– 421. Crawford, R. P., J. D. Huber, and B. S. Adams. 1990. Epidemiology and surveillance. In: K. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 131–151. Crawford, R. M., L. L. van der Verg, L. Yuan, T. L. Hadfield, R. L. Warren, E. S. Drazek, H. H. Houng, C. Hammack, K. Sasala, T. Polsinelli, J. Thompson, and D. L. Hoover. 1996. Deletion of purE attenuates Brucella melitensis infection in mice. Infect. Immun. 64:2188–2192. Crespo-Leó n, F. 1994. Brucelosis ovina y caprina. Office International des Epizooties. Paris , France. 1–451. Cunningham, B. 1977a. Experiences with strain 45/20 vaccines in ireland. In: R. P. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 188–200. Cunningham, B. 1977b. Protective effects of calostral antibodies by B. abortus on strain 19 vaccination and field infection. Vet. Rec. 101:521–529. Da Costa, M., J. P. Guillou, B. Garin-Bastuji, M. Thiebaud, and G. Dubray. 1996. Specificity of six gene sequences for the detection of the genus Brucella by DNA amplification. J. Appl. Bacteriol. 81:267–275. Dalrymple-Champneys, W. 1960. Brucella Infection and Undulant Fever in Man. Oxford University Press. London, UK. 95–114. Dasinger, B. L., and J. B. Wilson. 1962. Glutamate metabolism in Brucella abortus strains of low and high virulence. J. Bacteriol. 84:911–915.

CHAPTER 3.1.16 Davis, D. S. 1990. Brucellosis in wild life. In: K. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 321–334. Dazord, L., Y. Le Garrec, M. Bonnier, and L. Toujas. 1980. Increased resistance to tumor graft in mice infected by vaccinal strains of Brucella abortus. Recent Results Cancer Res. 75:92–99. Dazord, L., A. Martin, R. Le Rest, P. Y. Le Prise, J. J. Larzul, C. Gandhour, C. David, and L. Toujas. 1984. Phase I study of immunotherapy by live Brucella abortus (strain 19 BA) in cancer patients. Cancer Treat. Rep. 68:417– 418. De Bolpe, J., L. D., and C. Garcia-Carrillo. 1991. Vacuna fenol-insoluble contra la brucelosis humana: Evaluacion del poder inmunogenico en cobayos. Rev. Inst. Med. Trop. São Paulo 33:23–27. Dees, C., and R. D. Schultz. 1990. The mechanism of enhanced intraphagocytic killing of bacteria by liposomes containing antibiotics. Vet. Immunol. Immunopathol. 24:135–146. De Fays K., A. Tibor, C. Lambert, C. Vinals, P. Denoel, X. de Bolle, J. Wouters, J. J. Letesson, and E. Depiereux. 1999. Structure and function prediction of the Brucella abortus P39 protein by comparative modeling with marginal sequence similarities. Protein. Eng. 12:217–223. Dehio, C., and M. Meyer. 1997. Maintenance of broad-range incompatibility group P and group Q plasmids and transposition of Tn5 in Bartonella henselae following conjugal plasmid transfer from Escherichia coli. J. Bacteriol. 179:538–540. De Lajudie, P., A. Willems, B. Pot, D. Dewettinck, G. Maestrojuan, M. Neyra, M. D. Collins, B. Dreyfus, K. Kersters, and M. Gillis. 1994. Polyphasic taxonomy of rhizobia: Emendation of the genus Shinorizobium and description of Sinorhizobium meliloti comb. nov., Shinorizobium saheli sp. nov., and Sinorhizobium teranga sp. nov. Int. J. Syst. Bacteriol. 44:715–733. De Ley, J., W. Mannheim, P. Segers, A. Livens, M. Denijin, M. Vanhoucke, and M. Gills. 1987. Ribosomal ribonucleic acid cistron similarities and taxonomic neighborhood of Brucella and CDC Group Vd. Int. J. Syst. Bacteriol. 37:35–42. Delrue, R. M., M. Martinez-Lorenzo, P. Lestrate; I. Danese; V. Bielarz, P. Mertens, X. DeBolle, A. Tibor; J. P. Gorvel, and J. J. Letesson. 2001. Identificatoin of Brucellas genes involved in intracellular trafficking. Cell. Microbiol. 3:487–497. DelVecchio, V. G., V. Kapatral, R. J. Redkar, G. Patra, C. Mujer, T. Los, N. Ivanova, I. Anderson, A. Bhattacharyya, A. Lykidis, G. Reznik, L. Jablonski, N. Larsen, M. D’Souza, A. Bernal, M. Mazur, E. Goltsman, E. Selkov, P. Elzer, S. Hagius, D. O’Callaghan, J. J. Letesson, R. Haselkorn, N. Kyrpides, and R. Overbeek. 2002. The genome sequence of the facultative intracellular pathogen Brucella melitensis. Proc. Natl. Acad. Sci. USA 99:443–448. D’Enfert, C., L. J. Wuestehube, T. Lila, and R. Schekman. 1991. Sec12β-dependent membrane binding of the small GTP-binding protein Sarpl promotes formation of transport vesicles from the endoplasmic reticulum. J. Cell Biol. 114:663–670. Denoel, P. A., M. S. Zygmunt, V. Weynants, A. Tibor, B. Lichtfouse, P. Briffeuil, J. N. Limet, and J. J. Letesson. 1995. Cloning and sequencing of the bacterioferritin gene of Brucella melitensis 16M strain. FEBS Lett. 361:238–242.

The Genus Brucella

437

Denoel, P. A., R. M. Crawford, M. S. Zygmunt, A. Tibor, V. E. Weynants, F. Godfroid, D. L. Hoover, and J. J. Letesson. 1997a. Survival of bacterioferritin deletion mutant of Brucella melitensis 16M in human monocytederived macrophages. Infect. Immun. 65:4337–4340. Denoel, P. A., T. K. Vo, A. Tibor, V. E. Weynants, J. M. Trunde, G. Dubray, J. N. Limet, and J. J. Letesson. 1997b. Characterization, occurrence, and molecular cloning of a 39kilodalton Brucella abortus cytoplasmic protein immunodominant in cattle. Infect. Immun. 65:495–502. De Petris, S., G. Karlsbad, and R. W. I. Kessel. 1964. The ultrastructure of S and R variants of Brucella abortus grown on a lifless medium. J. Gen. Microbiol. 35:373–382. De Siervo, A. J., and A. D. Homola. 1980. Analysis of Caulobacter crescentus lipids. J. Bacteriol. 143:1215–1222. Desjardins, M., L. A. Huber, R. G. Parton, and G. Griffiths. 1994. Biogenesis of phagolysosomes proceeds though a sequential series of interactions with the endocytic apparatus. J. Cell Biol. 124:677–688. Desjardins, M., and A. Descoteaux. 1997. Inhibition of phagolysosomal biogenesis by Leishmania lipophosphoglycan. J. Exp. Med. 185:2061–2068. Detilleux, P. G., N. F. Cheville, and B. L. Deyoe. 1988. Pathogenesis of Brucella abortus in chicken embryos. Vet. Pathol. 25:138–146. Detilleux, P. G., B. L. Deyoe, and N. F. Cheville. 1990a. Entry and intracellular localization of Brucella spp. in Vero cells: Fluorescence and electron microscopy. Vet. Pathol. 27:317–328. Detilleux, P. G., B. L. Deyoe, and N. F. Cheville. 1990b. Penetration and intracellular growth of Brucella abortus in non-phagocytic cells in vitro. Infect. Immun 58:2320– 2328. Detilleux, P. G., B. L. Deyoe, and N. F. Cheville. 1991. Effect on endocytic and metabolic inhibitors on the internalization and intracellular growth of Brucella abortus in Vero cells. Amer. J. Vet. Res. 52:1658–1664. Díaz, R., and N. Bosseray. 1973a. Identification d’un composé antigenique specifique de la phase rugueuse des Brucella. Ann. Rech.Vet. 4:283–292. Díaz, R., and L. M. Jones. 1973b. The immuno-diffusion method for the identification of cattle vaccinated with Brucella abortus strain 45-20. Vet. Rec. 93:300–302. Díaz, R., and N. Bosseray. 1974. Estudio de las relaciones antigénicas entre Yersinia enterocolitica serotipo 9 y otras especies bacterianas Gram-negativas. Microbiology 27:1–14. Díaz, R. I., E. Maravi-Poma, and A. Rivero. 1976. Comparison of counter-immunoelectropjhoresis with other serological tests in the diagnosis of human brucellosis. Bull. WHO 53:417–124. Díaz, R., and M. A. Oyeledum. 1977. Studies of some biological activities of Brucella endotoxin in normal and infected animals and the role of hypersensitivity factor. Ann. Sclavo 19:117–130. Díaz, R., P. Garatea, L. M. Jones, and I. Moriyó n. 1979. Radial immunodiffusion test with a Brucella polysaccharide antigen for differentiating infected from vaccinated cattle. J. Clin. Microbiol. 10:37–41. Díaz, R., J. Toyos, M. D. Salvo, and M. L. Pardo. 1981. A simple method for the extraction of polysaccharide B from Brucella cells for use in the radial immunodiffusion test diagnosis of bovine brucellosis. Ann. Rech. Vet. 12:35–39. Díaz, R., and I. Moriyó n. 1989. Laboratory techniques in the diagnosis of human brucellosis. In: E. J. Young and M.

438

E. Moreno and I. Moriyó n

J. Corbel (Eds.) Brucellosis: Clinical and Laboratory Aspects of Human Infection. CRC Press. Boca Raton, FL. 73–83. Díaz-Aparicio, E., V. Aragó n, C. Marín, B. Alonso, M. Font, E. Moreno, S. Pérez-Ortiz, J.-M. Blasco, R. Díaz, I. Mroiyó n. 1993. Comparative analysis of Brucella serotype A and M Yersinia enterocolitica O:9 polysaccharides for serological diagnosis of brucellosis in cattle, sheep and goats. J. Clin. Microbiol. 31:3136–3141. Díaz-Aparicio, E., C. M. Marín, B. Alonso-Urmeneta, V. Aragó n, S. Pérez-Gó mez, M. L. Pardo, J. M. Blasco, R. Díaz, and I. Moriyó n. 1994. Evaluation of serological tests for diagnosis of Brucella melitensis infection of goats. J. Clin. Microbiol. 32:1159–1165. Dickerson, R. E. 1980. Cytochrome c and the evolution of energy metabolism. Sci. Amer. 242:136–153. Doborowski, J. M., and L. D. Sibley. 1996. Toxoplasma invasion of mammalian cells is powered by the actin cytoskeleton. Cell. 84:933–939. Dobzhansky, T. 1937. Genetics and the Origin of Species. Columbia University Press. New York, NY. 1–457. Dorrell, N., S. Spencer, V. Foulonge, P. Guigue-Talet, D. O’Callaghan, and B. W. Wren. 1998. Identification, cloning and initial characterisation of FeuPQ in Brucella suis: A new sub-family of two-component regulatory systems. FEMS Microbiol. Lett. 162:143–150. Dorrell, N., P. Guigue-Talet, S. Spencer, V. Foulonge, D. O’Callaghan, and B. W. Wren. 1999. Investigation into the role of the response regulator NtrC in the metabolism and virulence of Brucella suis. Microb. Pathog. 27:1–11. Dorsh, M., E. Moreno, and E. Stackebrandt. 1989. Nucleotide sequence of 16S rRNA from Brucella abortus. Nucl. Acids Res. 17:1765. Douglas, J. T., E. Y. Rosenberg, H. Nikaido, D. R. Verstreate, and A. J. Winter. 1984. Porins of Brucella species. Infect. Immun. 44:16–21. Douglas, J. T., and D. A. Palmer. 1988. Use of monoclonal antibodies to identify the distribution of A and M epitopes on smooth Brucella species. J. Clin. Microbiol. 26:1353–1356. Drancourt, M., P. Brouqui, and D. Raoult. 1997. Afipia clevelandensis antibodies and cross-reactivity with Brucella spp. and Yersinia enterocolitica O:9. Clin. Diagn. Lab. Immunol. 4:748–752. Drazek, E. S., H. S. Houng, R. M. Crawford, T. L. Hadfield, D. L. Hoover, and R. L. Warren. 1995. Deletion of purE attenuates Brucella melitensis 16M for growth in human monocyte-derived macrophages. Infect. Immun. 63:3297–3301. Dubray, G. 1972. Etude ultrastructurale des bacteries de colonies lisses (S) et rugueuses (R) du genre “Brucella.” Ann. Inst. Pasteur Paris 123:171–193. Dubray, G. 1975. Influence de la fixation sur la mise en évidence cytochimique des polysaccharides des bactéries du genre Brucella. J. Microsci. Biol. Cell. 24:59–74. Dubray, G. 1976. Localisation cellulaire des polyosides des bactéries des genres Brucella et Escherichia en phase lisse (S) ou rugueuse (R). Ann. Microbiol. 127:133– 149. Dubray, G. 1984. Progrès récents sur la biochimie et le propriétés biologiques des antigénes de Brucella. Dev. Biol. Stand. 56:131–150. Dunn, W. A. 1994. Autophagy and related mechanisms of lysosome-mediated protein degradation. Trends Cell Biol. 4:139–143.

CHAPTER 3.1.16 Eberhard, W. G. 1990. Evolution in bacterial plamids and levels of selection. Quarterly Rev. Biol. 65:3–22. Edmonds, M. D., G. G. Schurig, L. E. Samartino, P. G. Hoyt, J. V. Walker, S. D. Hagius, and P. H. Elzer. 1999. Biosafety of Brucella abortus strain RB51 for vaccination of mature bulls and pregnant heifers. Amer. J. Vet. Res. 60:722–725. Ekaza, E., L. Gulloteau, J. Teyssier, J. P. Liautard, and S. Koheler. 2000. Functional analysis of the ClpATPase ClpA of Brucella suis, and persistence of knockout mutant in BALB/c mice. Microbiology 46:1605–1616. Elberg, S. S., and D. J. Ralston. 1980. Enhancement of murine bone marrow colony formation and L-transformation by Brucella antigen. Can. J. Comp. Med. 44:320–327. Eldredge, N., and S. J. Gould. 1972. Punctuated equilibria: An alternative to phyletic gradualism. In: T. J. M. Schopf (Ed.) Models in Paleobiology. Freeman Cooper. San Francisco, CA. 82–115. el-Idrissi, A. H., A. Benkirane, M. el-Maadoudi, M. Bouslikhane, J. Berrada, and A. Zerouali. 2001. Comparison of the efficacy of Brucella abortus strain RB51 and Brucella melitensis Rev. 1 live vaccines against experimental infection with Brucella melitensis in pregnant ewes. Rev. Sci Tech. 20:741–747. Elzer, P. H., R. W. Phillips, M. E. Kovach, K. M. Peterson, and R. M. Roop. 1994. Characterization and genetic complementation of a Brucella abortus hightemperature-requirement A (htrA) deletion mutant. Infect. Immun. 62:4135–4139. Elzer, P. H., R. W. Phillips, G. T. Robertoson, and R. M. Roop 2nd. 1996. The HtrA stress response protease contributes to resistance of Brucella abortus to killing by murine phagocytes. Infect. Immun. 64:4838–4841. Elzer, P. H., F. M. Enright, J. R. McQuiston, S. M. Boyle, and G. G. Schurig. 1998. Evaluation of a rough mutant of Brucella melitensis in pregnant goats. Res. Vet. Sci. 64:259–260. Elzer, P., M. Edmonds, and A. Cloeckaert. 2000. Brucella mutants lacking the outer membrane protein Omp25. In: D. O’Callaghan (Ed.) Brucellosis 2000. INSERM, Facultéde Médicine. Nîmes, France. 77:79–80. Endley, S., D. McMurray, and T. A. Fich. 2001. Interruption of cydB locus in Brucella abortus attenuates intracellular survival and virulence in mouse model of infection. J. Bacteriol. 183 :2454–2462. Enright, F. M. 1990a. Mechanisms of self cure in Brucella abortus-infected cattle. In: L. G. Adams (Ed.) Advances in Brucellosis Research. Texas A&M University Press. College Station, TX. 191–196. Enright, F. M. 1990b. The pathogenesis and pathobiology of Brucella infection in domestic animals. Antigens of Brucella. In: K. Nielsen and B. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 301–320. Enright, F. M., and L. Samartino. 1994. Mechanisms of abortion in Brucella abortus infected cattle. Proceedings of the 98 annual meeting of the United States Animal Health Association, USDA. Richmond, Virginia. 88–95. Essenberg, R. C. 1995. Cloning and characterization of the glucokinase gene of Brucella abortus 19 and identification of three other genes. J. Bacteriol. 177:6297– 6300. Essenberg, R. C., C. Candler, and S. K. Nida. 1997. Brucella abortus strain 2308 putative glucose and galactose transporter gene: Cloning and characterization. Microbiology 143:1549–1555.

CHAPTER 3.1.16 Etemadi, H., A. Raissadat, M. J. Pickett, Y. Zafari, and P. Vahedifar. 1984. Isolation of Brucella spp. from clinical specimens. J. Clin. Microbiol. 20:586. Ewalt, D. R., R. A. Packer, and M. W. Harrison. 1983. An improved selective medium for isolating Brucella sp. from bovine milk. Proc. Int. Symp. Vet. Lab. Diagn. 3:577–589. Ewalt, D. R., J. B. Payeur, B. M. Martin, D. R. Cummins, and W. G. Miller. 1994. Characteristics of a Brucella species from a bottlenose dolphin (Tursiops truncatus). J. Vet. Diagn. Invest. 6:448–452. Ewalt, D. R., and B. J. Bricker. 2000. Validation of the abbreviated Brucella AMOS PCR as a rapid screening method for differentiation of Brucella abortus field strain isolates and the vaccine strains, 19 and RB51. J. Clin. Microbiol. 38:3085–3086. Eze, M. O., L. Yuan, R. M. Crawford, C. M. Paranavitana, T. L. Hadfield, A. K. Bhattacharjee, R. L. Warren, and D. L. Hoover. 2000. Effects of opsonization and gamma interferon on growth of Brucella melitensis 16M in mouse peritoneal macrophages in vitro. Infect. Immun. 68:257–263. FAO/WHO. 1986. Joint Report, Expert Committee on Brucellosis: World Health Organization Technical Report. FAO/WHO Publications. Geneva , Switzerland. 1–77. Farrell, I. D. 1974. The development of a new selective medium for the isolation of Brucella abortus from contaminated sources. Res. Vet. Sci. 16:280–286. Fekete, A., J. A. Bantle, and S. M. Halling. 1992a. Detection of Brucella by polymerase chain reaction in bovine fetal and maternal tissues. J. Vet. Diagn. Invest. 4:79–83. Fekete, A., J. A. Bantle, S. M. Halling, and R. W. Stich. 1992b. Amplification fragment length polymorphism in Brucella strains by use of polymerase chain reaction with arbitrary primers. J. Bacteriol. 174:7778–7783. Fensterbank, R., J. M. Verger, and M. Grayon. 1987. Conjunctival vaccination of young goats with Brucella melitensis strain Rev. 1. Ann. Rech. Vet. 18:397–403. Fernandez-Lago, L., M. Monte, and A. Chordi. 1996. Endogenous gamma interferon and interleukin−10 in Brucella abortus 2308 infected mice. FEMS Immunol. Med. Microbiol. 15:109–114. Ferrari, G., H. Langen, M. Naito, and J. Pieters. 1999. A coat protein on phagosomes involved in the intracellular survival of Mycobacteria. Cell 97:435–447. Ficapal, A., B. Alonso-Urmeneta, J. Velasco, I. Moriyó n, and J. M. Blasco. 1995. Diagnosis of Brucella ovis infection of rams with an ELISA using protein G as conjugate. Vet. Rec. 137:145–147. Ficht, T. A., S. W. Bearden, B. A. Sowa, and L. G. Adams. 1989. DNA sequence and expression of the 36-kilodalton outer membrane protein gene of Brucella abortus. Infect. Immun. 57:3281–3291. Ficht, T. A., S. W. Bearden, B. A. Sowa, and H. Marquis. 1990. Genetic variation at the omp2 porin locus of the brucellae: Species-specific markers. Molec. Microbiol. 4:1135– 1142. Ficht, T. A., H. A. Husseinen, J. Derr, and W. Bearden. 1996. Species specific sequences at the omp2 locus for Brucella type strains. Int. J. Syst. Bacteriol. 46:329–331. Finlay, B., and P. Cossart. 1997a. Exploitation of mammalian host cell functions by bacterial pathogens. Science 276:934–938. Finlay, B., and S. Falkow. 1997b. Common themes in microbial pathogenicity revised. Microbiol. Molec. Biol. Rev. 61:136–169.

The Genus Brucella

439

Flesh, I. E. A., J. H. Hess, I. P. Oswald, and S. H. E. Kaufmann. 1994. Growth inhibition of Mycobacterium bovis by IFN-alpha stimulated macrophages: Regulation by endogenous tumor necrosis factor-alpha and by IL10. Int. Immunol. 6:693–700. Flores-Castro, R., and G. M. Baer. 1979. Brucellosis (Brucella melitensis)— Zoonotic implications. In: J. H. Steele (Ed.) Handbook Series in Zoonoses. CRC Press. Boca Raton, FL. 195–225. Flowers, A. I. 1977. National programs. In: R. P. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 333–415. Flynn, J. L., M. M. Goldstein, J. Chan, K. J. Triebold, K. Pfeffer, C. J. Lowenstein, R. Schreiberg, T. W. Mak, and B. R. Bloom. 1995. Tumor necrosis factor-alpha is required in the protective immune response against Mycobacterium tuberculosis in mice. Immunity 2:561– 572. Fonstein, M., S. Zheng, and R. Haselkorn. 1992. Physical map and genome of Rhodobacter capsulatus SB1003. J. Bacteriol. 174:4070–4077. Forestier, C., E. Moreno, S. Méresse, A. Phalipon, J. PizarroCerdá, D. Olive, P. Sansonetti, and J. P. Gorvel. 1999a. Interaction of Brucella abortus lipopolysaccharide with major histocompatibility complex class-II molecules in B lymphocytes. Infect. Immun. 67:4048–4054. Forestier, C., E. Moreno, J. Pizarro-Cerdá, P. Sansonetti, and J. P. Gorvel. 1999b. Lysosomal accumulation and recycling of LPS to cell surface of murine macrophages, an in vitro and in vivo study. J. Immunol. 162:6784–6791. Forestier, C., F. Deleuil, N. Lapaque, E. Moreno, and J. P. Gorvel. 2000. Brucella abortus lipopolysaccharide in murine peritoneal macrophages acts as a down-regulator of T cell activation. J. Immunol. 165:5202–5210. Forschner, E., I. Bunger, H. P. Krause, and D. Kuttler. 1989. Kontrollmassnahmen in amtlich anerkannten brucellosefreien und leukoseunverdachtigen Milchviehbestanden auf der Basis von Tankmilch-Proben in Kombination mit ELISA-Tests. Deutsch. Tierarztl. Wochenschr. 96:475–486. Foulongne, V., G. Bourg, C. Cazevieille, A. MichauxCharachon, and D. O’Callaghan. 2000. Identification of Brucella suis genes affecting intracellular survival in an in vitro human macrophage infection model by signature-tagged transposon mutagenesis. Infect. Immun. 68:1297–1303. Foulongne, V., K. Walravens, G. Bourg, M. L. Boschiroli, J. Godofroid, M. Ramuz, D. O’Callaghan. 2001. Aromatic compound-dependent Brucella suis is attenuated in both cultured cells and mouse models. Infect. Immun. 69:547– 550. Fox, G. E., E. Stackebrandt, R. B. Hespell, J. Gibson, J. Maniloff, T. A. Dyer, R. S. Wolfe, W. E. Balch, R. S. Tanner, L. J. Magrum, L. B. Zablen, R. Blakemore, R. Gupta, L. Bonen, B. J. Lewis, D.-A. Stahl, K. R. Luehrsen, K. N. Chen, and C. R. Woese. 1980. The phylogeny of prokaryotes. Science 209:457–463. Foz, A., L. Arcalis, S. Garriga, J. Manzanares, and F. Ortiz. 1954. Valor de algunos métodos de aboratorio en el diagnó stico de la brucelosis humana. Rev. Diag. Biol. 3:349–383. Frank, M. M. 1992. The mechanism by which microorganisms avoid complement attack. Curr. Opin. Immunol. 4:4–19. Frank, S. A. 1994. Kin selection and virulence in the evolution of protocells and parasites. Proc. Royal Soc. Lond. Series B, Biol. Sci. 258:153–161.

440

E. Moreno and I. Moriyó n

Freer, E., N. Rojas, A. Weintraub, A. Lindberg, and E. Moreno. 1995. Heterogeneity of Brucella abortus lipopolysaccharides. Res. Microbiol. 146:569–578. Freer, E., E. Moreno, I. Moriyó n, J. Pizarro-Cerdá, A. Weintraub, and J. P. Gorvel. 1996. Brucella/Salmonellalipopolysaccharide chimeras are less permeable to hydrophobic probes and more sensitive to cationic peptides and EDTA than their native Brucella spp. counterparts. J. Bacteriol. 178:5867–5876. Freer, E., J. Pizarro-Cerdá, A. Weintraub, J.-A. Bengoechea, I. Moriyó n, K. Hultemby, J. P. Gorvel, and E. Moreno. 1999. The outer membrane of Brucella ovis shows increased permeability to hydrophobic probes and is more susceptible to cationic peptides than are the outer membranes of mutant rough Brucella abortus strains. Infect. Immun. 67:6181–6186. Frenchick, P. J., R. J. F. Markam, A. H. Cochrane. 1985. Inhibition of phagosome-lysosome fusion in macrophages by soluble extracts of virulent Brucella abortus. Amer. J. Vet. Res. 46:332–335. Fumarola, D., S. Pece, R. Fumarulo, R. Petruzzelli, B. Greco, G. Giuliani, A. B. Maffione, and E. Jirillo. 1994. Downregulation of human polymorphonuclear cell activities exerted by microorganisms belonging to the alpha-2 subgroup of Proteobacteria (Afipia felis and Rochalimaea henselea). Immunopharma. Immunotoxi. 16:449– 461. Galdiero, M., C. Bentivoglio, I. Nuzzo, L. DeMartino, M. Molitierno, and C. R. Carratelli. 1995. Immunological response of mice after long-term stimulation with cell wall antigen from Brucella melitensis. Res. Microbiol. 146:507–515. Galibert F., T. M. Finan, S. R. Long, A. Puhler, P. Abola, F. Ampe, F. Barloy-Hubler, M. J. Barnett, A. Becker, P. Boistard, G. Bothe, M. Boutry, L. Bowser, J. Buhrmester, E. Cadieu, D. Capela, P. Chain, A. Cowie, R. W. Davis, S. Dreano, N. A. Federspiel, R. F. Fisher, S. Gloux, T. Godrie, A. Goffeau, B. Golding, J. Gouzy, M. Gurjal, I. Hernandez-Lucas, A. Hong, L. Huizar, R. W. Hyman, T. Jones, D. Kahn, M. L. Kahn, S. Kalman, D. H. Keating, E. Kiss, C. Komp, V. Lelaure, D. Masuy, C. Palm, M. C. Peck, T. M. Pohl, D. Portetelle, B. Purnelle, U. Ramsperger, R. Surzycki, P. Thebault, M. Vandenbol, F. J. Vorholter, S. Weidner, D. H. Wells, K. Wong, K. C. Yeh, and J. Batut. 2001. The composite genome of the legume symbiont Sinorhizobium meliloti. Science 293:668–672. Gall, D., and K. Nielsen. 1994. Improvements to the competitive ELISA for detection of antibodies to Brucella abortus in cattle sera. J. Immunoassay. 15:277–291. Gall, D. E., A. Colling, O. Maríno, E. Moreno, K. H. Nielsen, B. Perez, and L. Samartino. 1998. Enzyme immunoassays for serological diagnosis of bovine brucellosis: A trial in Latin America. Clin. Diagn. Lab. Immunol. 5:654–661. Gamazo, C., and I. Moriyó n. 1987. Release of outer membrane fragments by exponentially growing Brucella melitensis cells. Infect. Immun. 55:609–615. Gamazo, C., A. J. Winter, I. Moriyó n, J. I. Riezu-Boj, J. M. Blasco, and R. Díaz. 1989. Comparative analyses of proteins extracted by hot saline or released spontaneously into outer membrane blebs from field strains of Brucella ovis and Brucella melitensis. Infect. Immun. 57:1419– 1426. García-Carrillo, C. 1980. Comparison of B. melitensis Rev. 1 and B. abortus strain 19 as a vaccine against brucellosis in cattle. Zentralbl. Veterinarmed. B 27:131–137.

CHAPTER 3.1.16 García-Carrillo, C. 1990. Laboratory animal models for brucellosis studies. In: K. Nielsen, and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 423– 442. Gargani, G. 1977. Attempts at a system of numerical taxonomy of the genus Brucella. Ann. Scalvo 19:61–66. Garin, J., R. Diez, S. Kieffer, J. F. Dermine, S. Duclos, E. Gagnon, R. Sadoul, C. Rondeau, M. Desjardins. 2001. The phagosome proteome: Insight into phagosome functions. J. Cell Biol. 152:165–80. Garin-Bastuji, B., J. M. Blasco, M. Grayon, and J. M. Verger. 1998. Brucella melitensis infection in sheep: Present and future. Vet. Res. 29:255–274. Garin-Bastuji, B., N. Hummel, G. Gerbier, C. Cau, R. Pouillot, M. Da Costa, and J. J. Fontaine. 1999. Non specific serological reactions in the diagnosis of bovine brucellosis: Experimental oral infection of cattle with repeated doses of Yersinia enterocolitica O:9. Vet. Microbiol. 66:223–233. Garner, M. M., D. M. Lambourn, S. J. Jeffries, B. J. Hall, J. C. Rhyan, D. R. Ewalt, L. M. Polzin, and N. F. Cheville. 1997. Evidence of Brucella infection in Parafilaroides lungworms in a pacific harbor seal (Phoca vitulina richardsi). J. Vet. Diagn. Invest. 9:298–303. Gaviria-Ruiz, M. M., and N. M. Cardona-Castro. 1995. Evaluation and comparison of different blood culture techniques for bacteriological isolation of Salmonella typhi and Brucella abortus. J. Clin. Microbiol. 33:868–871. Gay, B., H. Mauss, and S. Sanchez-Teff. 1981. Aspect ultrastructuraux de la phagocytose in vivo et in vitro de Brucella par les macrophages du péritoine de la souris. Ann. Immunol. 132 D:299–313. Gay, B., H. Mauss, and S. Sanchez-Teff. 1986. Identification of fibronectins in peritoneal macrophages during the phagocytosis of Brucella: An immunocytochemical study by electron microscopy. Virchows Arch. B: Cell Pathol. 52:169–176. Gerhardt, P., H. B. Levine, and J. B. Wilson. 1950a. The oxidative dissimilation of amino acids and related compounds by Brucella abortus. J. Bacteriol. 60:459–467. Gerhardt, P., L. A. Tucker, and J. B. Wilson. 1950b. The nutrition of Brucellae: Utilization of single amino acids for growth. J. Bacteriol. 59:777–782. Gerhardt, P. 1958. The nutrition of Brucellae. Bacteriol. Rev. 22:81–98. Giauffret, A., and R. Sanchis. 1974. Etude d’un foyer d’epididymite contagieuse du bélier: Eradication de la maladie. Bull. Off. Int. Epizoot. 82:581–589. Gibby, I. W., and A. M. Gibby. 1965. Host-parasite relationships with Brucella neotomae. J. Bacteriol. 80:9–16. Godfroid, F., B. Taminiau, I. Danese, P. Denoel, A. Tibor, V. Weynants, A. Cloeckaert, J. Godfroid, and J. J. Letesson. 1998. Identification of the perosamine synthetase gene of Brucella melitensis 16M and involvement of lipopolysaccharide O side chain in Brucella survival in mice and in macrophages. Infect. Immun. 66:5485–5493. Godfroid, F., A. Cloeckaert, B. Taminiau, I. Danese, A. Tibor, X. de Bolle, P. Mertens, and J. J. Letesson. 2000. Genetic organisation of the lipopolysaccharide O-antigen biosynthesis region of Brucella melitensis 16M (wbk). Res. Microbiol. 151:655–668. Goldbaum, F. A., C. P. Rubbi, J. C. Wallach, S. E. Miguel, P. C. Baldi, and C. A. Fossati. 1992. Differentiation between active and inactive human brucellosis by measuring antiprotein humoral immune responses. J. Clin. Microbiol. 30:604–607.

CHAPTER 3.1.16 Goldbaum, F. A., C. A. Velikovsky, P. C. Baldi, S. Mortl, A. Bacher, and C. A. Fossati. 1999. The 18-kDa cytoplasmic protein of Brucella species. An antigen useful for diagnosis is a lumazine synthase. J. Med. Microbiol. 48:833– 839. Golding, B. J. I., P. Highet, R. Blackburn, J. Manischewitz, N. Blyveis, D. Agus, and H. Golding. 1995. Brucella abortus conjugated with a gp120 or V3 loop peptide derived from human immunodeficient virus (HIV) type 1 induces neutralizing anti-HIV antibodies, and the V-3 B. abortus conjugate is effective even after CD4 T-cell depletion. J. Virol. 69:3299–3307. Goldstein, J., T. Hoffman, C. Frasch, E. F. Lizzio, P. R. Beining, D. Hochstein, Y. L. Lee, R. D. Angus, and B. Golding. 1992. Lipopolysaccharide (LPS) from Brucella abortus is less toxic than that from Escherichia coli, suggesting the possible use of B. abortus or LPS from B. abortus as carrier in vaccines. Infect. Immun. 60:1385– 1389. Gó mez-Miguel, M. J., and I. Moriyó n. 1986. Demonstration of a peptidoglycan-linked lipoprotein and characterization of its trypsin fragment in the outer membrane of Brucella spp. Infect. Immun. 53:678–684. González-Carrero, M. I., F. J. Sangari, J. Aguero, J. M. García-Lobo. 2002. Brucella abortus strain 2308 produces brucebactin, a highly efficient catecholic siderophore. Microbiology 148:353–360. Goodner B., G. Hinkle, S. Gattung, N. Miller, M. Blanchard, B. Qurollo, B. S. Goldman, Y. Cao, M. Askenazi, C. Halling, L. Mullin, K. Houmiel, J. Gordon, M. Vaudin, O. Iartchouk, A. Epp, F. Liu, C. Wollam, M. Allinger, D. Doughty, C. Scott, C. Lappas, B. Markelz, C. Flanagan, C. Crowell, J. Gurson, C. Lomo, C. Sear, G. Strub, C. Cielo, and S. Slater. 2001. Genome sequence of the plant pathogen and biotechnology agent Agrobacterium tumefaciens C58. Science 294:2323–2328. Graham, P. H., M. J. Sadowsky, H. H. Keyser, Y. M. Barnet, R. S. Bradley, J. E. Cooper, D. J. De Ley, B. D. W. Jarvis, E. B. Roslycky, B. W. Strijdom, and J. P. W. Young. 1984. Proposed minimal standards for the description of new genera and species of root- and steam-nodulating bacteria. Int. J. Syst. Bacteriol. 41:582–587. Granfors, K., M. K. Viljanen, and A. Toivanen. 1981. Measurement of immunoglobulin M, immunoglobulin G, and immunoglobulin A antibodies against Yersinia enterocolitica by enzyme-linked immunosorbent assay: Comparison of lipopolysaccharide and whole bacterium as antigen. J. Clin. Microbiol. 14:6–14. Grilló , M. J. 1997. Brucelosis experimental en ratones: Control de calidad de vacunas y estudio de factores de virulencia en Brucella (PhD thesis). University of Zaragoza. Zaragoza, Spain. Grilló , M. J., N. Bosseray, and J. M. Blasco. 2000. In vitro markers and biological activity in mice of seed lot strains and commercial Brucella melitensis Rev 1 and Brucella abortus B19 vaccines. Biologicals 28:119–127. Grimont, F., J. M. Verger, P. Cornelis, J. N. Limet, M. Lefevre, M. Grayon, B. Regnault, B. J. Van, and P. A. Grimont. 1992. Molecular typing of Brucella with cloned DNA probes. Res. Microbiol. 143:55–65. Gross, A., S. Spiesser, A. Terraza, B. Rouot, E. Caron, and J. Dornand. 1998. Expression and bactericidal activity of nitric oxide syntase in Brucella suis infected-murine macrophages. Infect. Immun. 66:1309–1316. Gross, A., A. Terraza, S. Ouahrani-Bettache, J. P. Liautard, and J. Dornand. 2000. In vitro Brucella suis infection

The Genus Brucella

441

prevents the programmed cell death of human monocytic cells. Infect. Immun. 68:342–351. Guerra, M. A., and P. Nicoletti. 1986. Comparison of the susceptibility of Brucella abortus isolates obtained before and after cows were treated with oxytetracycline and streptomycin. Amer. J. Vet. Res. 47:2612–2613. Gunn, J. S., and S. I. Miller. 1996. PhoP-PhoQ activates transcription of pmrAB, encoding a two-component regulatory system involved in Salmonella typhimurium antimicrobial peptide resistance. J. Bacteriol. 178:6857– 6864. Gupta, R. S. 1997. Protein phylogenies and signaturesevolutionary relationships within prokaryottes and between prokaryotes and eukaryotes. Ant. v. Leeuwenhoek 72:49–61. Guttman, D. S., and D. E. Dykhuizen. 1994. Clonal divergence in Escherichia coli as a result of recombination, not mutation. Science 266:1380–1382. Guzmán-Verri, C., E. Chaves-Olarte, C. von Eichel-Streiber, I. Ló pez-Goñ i, M. Thelestam, S. Arvidson, J-P. Gorvel, and E. Moreno. 2001. GTPases of the Rho subfamily are required for Brucella abortus internalization in nonprofessional phagocytes: direct activation of Cdc42. J. Biol. Chem. 276:44435–44443. Halliday, R. 1968. Effect of passive immunization against Brucella abortus on active production of Brucella abortus agglutinins in joung rats. J. Pathol. Bacteriol. 96:137– 146. Halling, S. M., and E. S. Zehr. 1990. Polymorphism in Brucella spp. due to highly repeated DNA. J. Bacteriol. 172:6637–6640. Halling, S. M., F. M. Tatum, and B. J. Bricker. 1993. Sequence and characterization of an insertion sequence, IS711, from Brucella ovis. Gene 133:123–127. Halling, S. M., and B. J. Bricker. 1994. Characterization and occurrence of two repeated palindromic DNA elements of Brucella spp. Bru-RS1 and Bru-RS2. Molec. Microbiol. 14:681–689. Harmon, B. G., and L. G. Adams. 1987. Assessment of bovine mammary gland macrophage oxidative burst activity in chemiluminescence assay. Amer. J. Vet. Res. 48:119–125. Harmon, B. G., L. G. Adams, and M. Frey. 1988. Survival of rough and smooth strains of Brucella abortus in bovine mammary gland macrophages. Amer. J. Vet. Res. 49:1092–1097. Harmon, B. G., L. G. Adams, J. W. Templeton, and R. Smith. 1989. Macrophage function in mammary glands of Brucella abortus-infected cows and cows that resisted infection after inoculation of Brucella abortus. Amer. J. Vet. Res. 50:459–465. Harricane, M. C., E. Caron, F. Porte, and J. P. Liautard. 1996. Distribution of annexin I during non-pathogen or pathogen phagocytosis by confocal imaging and immunogold electron microscopy. Cell Biol. Inter. 20:193–203. Hatten, B. A., and S. E. Sulkin. 1966a. Intracellular production of Brucella L forms. 1. Recovery of L forms from tissue culture cells infected with Brucella abortus. J. Bacteriol. 91:285–296. Hatten, B. A., and S. E. Sulkin. 1966b. Intracellular production of Brucella L forms II: Induction and survival of Brucella abortus L forms in tissue culture. J. Bacteriol. 91:14–20. Hatten, B. A., M. L. Schulze, S. Y. Huang, and S. E. Sulkin. 1969. Ultrastructure of Brucella abortus L-forms induced by penicillin in a liquid and in a semisolid medium. J. Bacteriol. 99:611–618.

442

E. Moreno and I. Moriyó n

Hayes, F. A. 1977. Wildlife reservoirs of brucellosis. In: R. P. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 269– 276. Heck, F. C., J. D. Williams, J. Pruett, R. Sanders, and D. L. Zink. 1980. Enzyme-linked immunosorbent assay for detecting antibodies to Brucella abortus in bovine milk and serum. Amer. J. Vet. Res. 41:2082–2084. Herman, L., and H. De Ridder. 1992. Identification of Brucella spp. by using the polymerase chain reaction. Appl. Environ. Microbiol. 58:2099–2101. Hidalgo, R. J. 1977. Immunization. In: R. P. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 175–265. Higgins, R. 2000. Bacteria and fungi of marine mammals: A review. Can. Vet. J. 41:105–116. Hoerlin, A. B. 1957. The influence of colostrum on antibody response in baby pigs. J. Immunol. 78:112–119. Hoffmann, E. M., and J. J. Houle. 1984. Failure of Brucella abortus lipopolysaccharide (LPS) to activate the alternative pathway. Vet. Immunol. Immunopathol. 5:65–68. Holland, J. J., and M. J. Pickett. 1956. Intracellular behavior of Brucella variants in chick embryo cells in tissue culture. Proc. Soc. Exp. Biol. Med. 93:476–479. Hollingdale, M. R., J. W. Vinson, and J. E. Herrmann. 1980. Immunochemical and biological properties of the outer membrane-associated lipopolysaccharide and protein of Rochalimaea quintana. J. Infect. Dis. 141:672–679. Holmes, B., M. Popoff, M. Kiredjian, and K. Kersters. 1988. Ochrobactrum anthropi gen. nov., sp. nov. from human clinical specimens and previously known as group Vd. Int. J. Syst. Bacteriol. 38:406–416. Honeycutt, R. J., M. McClelland, and B. W. S. Sobral. 1993. Physical map of the genome of Rhizobium meliloti 1021. J. Bacteriol. 175:6945–6952. Hong, P., R. Tsolis, and T. A. Ficht. 2000. Identification if genes for chronic persistence of Brucella abortus in mice. Infect. Immun. 68:4102–4107. Hoover, D., R. M. Crawford, L. L. van der Verg, M. J. Izadjoo, A. K. Bhattacharjee, C. M. Paranavitana, R. L. Warren, M. P. Nikolich, and T. L. Hadfield. 1999. Protection of mice against brucellosis by vaccination with Bucella melitensis WR201(16MδpurEK). Infect. Immun. 67:5877–5884. Horwell, F. D., and C. Van Drimmelen. 1971. Brucella melitensis as a vaccine for cattle. J. Afr. Vet. Med. Assoc. 42:233–241. Huang, L. Y., A. M. Krieg, N. Eller, and D. E. Scott. 1999. Induction and regulation of Th1.inducing cytokines by bacterial DNA, lipopolysaccharide, and heat-inactivated bacteria. Infect. Immun. 26:6257–6263. Huber, J. D., and P. Nicoletti. 1986. Comparison of the results of card, rivanol, complement-fixation, and milk ring tests with the isolation rate of Brucella abortus from cattle. Amer. J. Vet. Res. 47:1529–1531. Huber, J. D., V. C. Beal, R. P. Crawford, and L. G. Adams. 1990. An evaluation of reduced dose Brucella abortus strain 19 vaccination. In: L. G. Adams (Ed.) Advances in Brucellosis Research. Texas A&M University Press. College Station, TX. 228–249. Huber, I., and S. Selenskapobell. 1994. Pulse field electrophoresis-fingerprinting, genome size estimation and rrn loci number of Rhizobium galegae. J. Appl. Bacteriol. 77:528–533. Huddleson, I. F., and W. H. Stahl. 1943. Catalase activity of the species Brucella as a criterion of virulence: Studies

CHAPTER 3.1.16 in brucellosis II. Mich. State College Agr. Expt. Stat. Tech. Bull. 182:57–63. Huddleson, I. F. 1955. Effect of esters of long chain fatty acids in agar mediums on the growth of Brucella abortus in the presence of blood. Bacteriol. Proc. 122. Hulse, M., S. Johnson, and P. Ferrieri. 1993. Agrobacterium infections in humans: Experience at one hospital and review. Clin. Infect. Dis. 16:112–117. Hunziker, W., and H. J. Geuze. 1995. Intracellular trafficking of lysosomal membrane proteins. BioEssays 18:379– 388. Imhoff, J. F., and H. G. Trü per. 1984. Purple nonsulfur bacteria (Rhodospirillaceae) Pfenning and Trü per 171, 17AL. In: N. R. Krieg and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 1:1658. Iñ n de Iannino, N., G. Briones, M. Tolmasky, and A. R. ó Ugalde. 1998. Molecular characterization of cgs, the Brucella abortus cyclic β (1-2) glucan synthetase: Genetic complementation of Rhizobium meliloti ndvB and Agrobacterium tumefaciens chvB mutants. J. Bacteriol. 180:4392–4400. Iñ n de Iannino, N., G. Briones, F. Iannino, and R. A. Ugalde. ó 2000. Osmotic regulation of cyclic 1,2-beta-glucan synthesis. Microbiology 146:1735–1742. Issa, H., and M. Jamal. 2000. Brucellosis in children in south Jordan. East. Mediterr. Health J. 5:895–902. Isshiki, Y., Y. Haishima, S. Kondo, and K. Hisatsune. 1995. Immunochemistry of group A and Inaba C antigen factors constituting the O antigen of O1 Vibrio cholerae. Eur. J. Biochem. 229:583–588. Itoh, Y., B. Hemmer, R. Martin, and R. N. Germain. 1999. Serial TCR engagement and down-modulation by peptide: MHC molecule ligands: Relationship to the quality of individual TCR signaling events. J. Immunol. 162:2073–2080. Jahans, K. L., G. Foster, and E. S. Broughton. 1997. The characterization of Brucella strains isolated from marine mammals. Vet. Microbiol. 57:373–382. Jensen, A., E., N. F. Cheville, C. O. Thoen, A. P. MacMillan, and W. G. Miller. 1999. Genome fingerprinting and development of a dendrogram for Brucella spp. isolated from seals, porpoises and dolphins. J. Vet. Diagn. Invest. 11:152–157. Jiang, X., and C. L. Baldwin. 1993a. Effects of cytokines on the intracellular growth of Brucella abortus. Infect. Immun. 61:124–134. Jiang, X., and C. L. Baldwin. 1993b. Iron augments macrophage-mediated killing of Brucella abortus alone and in conjunction with interferon-gamma. Cell. Immunol. 148:397–407. Jiang, X., B. Leonard, R. Benson, and C. L. Baldwin. 1993c. Macrophage control of Brucella abortus by oxygen intermediates and nitric oxide. Cell. Immunol. 151:309– 319. Jiménez de Bagü és, M. P., C. M. Marín, M. Braberán, and J. M. Blasco. 1989. Responses of ewes to B. melitensis Rev1 vaccine administered by subcutaneous or conjunctival routes at different stages of pregnancy. Ann. Rech. Vet. 20:205–213. Jiménez de Bagü és, M. P., C. M. Marín, and J. M. Blasco. 1991. Effect of antibiotic therapy and strain-19 vaccination on the spread of Brucella melitensis within an infected dairy herd. Prev. Vet. Med. 11:17–24. Jiménez de Bagü és, M. P., C. M. Marín, J. M. Blasco, I. Moriyó n, and C. Gamazo. 1992. An ELISA with Bru-

CHAPTER 3.1.16 cella lipopolysaccharide antigen for the diagnosis of B. melitensis infection in sheep and for the evaluation of serological responses following subcutaneous or conjunctival B. melitensis strain Rev. 1 vaccination. Vet. Microbiol. 30:233–241. Jiménez de Bagü és, M. P., M. Braberán, C. M. Marín, and J. M. Blasco. 1995. The Brucella abortus RB51 vaccine does not confer protection against Brucella ovis in rams. Vaccine 13:301–304. Jones, L. M., V. Montgomery, and J. B. Wilson. 1965. Characteristics of carbon-dioxide-independent cultures of Brucella abortus isolated from cattle vaccinated with strain 19. J. Infect. Dis. 115:312–320. Jones, L. M., R. Díaz, and A. G. Taylor. 1973. Characterization of allergens prepared from smooth and rough strains of Brucella melitensis. Br. J. Exp. Pathol. 54:492– 508. Jones, L. M., D. T. Berman, E. Moreno, B. L. Deyoe, M. J. Gildorf, J. D. Huber, and P. Nicoletti. 1980. Evaluation of radial immunodiffussion test with polysaccharide B antigen for the diagnosis of bovine brucellosis. J. Clin. Microbiol. 12:753–760. Jones, S. M., and A. J. Winter. 1992. Survival of virulent and attenuated strains of Brucella abortus in normal and gamma interferon-activated murine peritoneal macrophages. Infect. Immun. 60:3011–3014. Jordan, D. C. 1984. Family III, Rhizobiaceae Conn 1938, 321AL. In: N. R. Krieg, and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. William and Wilkins. Baltimore, MD. 1:234–256. Jouen-Beades, F., E. Paris. C. Dieulois, J.-F. Lemeland, V. Barre-Dezelus, S. Marret, G. Humbert, J. Leroy, and F. Tron. 1997. In vivo and in vitro activation and expansion of γδ T cells aduring Listeria monocytogenes infection in humans. Infect. Immun. 65:4267. Jubier-Maurin, V., A. Rodrigue, S. Ouahrani-Bettache, M. Layssac, M. Mandran-Berthelot, S. Köhler, and J. P. Liautard. 2001. Identification of the nik gene cluster of Brucella suis: Regulation and contribution to urease activity. J. Bacteriol. 183:426–434. Jumas-Bilak, E., C. Maugard, S. Michaux-Charachon, A. Allardet-Servent, A. Perrin, D. O’Callaghan, and M. Ramuz. 1995. Insertion of a unique restriction site in the bacterial chromosomes for study of genome organization. Microbiology 141:2425–2432. Jumas-Bilak, E., S. Milchaux-Charachon, G. Bourg, D. O’Callaghan, and M. Ramuz. 1998a. Differences in Chromosome number and genome rearrangemnets in the genus Brucella. Molec. Microbiol. 27:99–106. Jumas-Bilak, E., S. Milchaux-Charachon, G. D. Bourg, M. Ramuz, and A. Allardet-Servent. 1998b. Unconventional genomic organization in the alpha subgroup of Proteobacteria. J. Bacteriol. 180:2749–2755. Kadowaki, M., R. Venerando, G. Miotto, and G. E. Mortimore. 1994. De novo autophagic vacuole formation in hematocytes permeabilized by Staphylococcus aureus alfa-toxin. J. Biol. Chem. 269:3703–3710. Kaneko T., Y. Nakamura, S. Sato, E. Asamizu, T. Kato, S. Sasamoto, A. Watanabe, K. Idesawa. A. Ishikawa, K. Kawashima, T. Kimura, Y. Kishida, C. Kiyokawa, M. Kohara, M. Matsumoto, A. Matsuno, Y. Mochizuki, S. Nakayama, N. Nakazaki, S. Shimpo, M. Sugimoto, C. Takeuchi, M. Yamada, and S. Tabata. 2000. Complete genome structure of the nitrogen-fixing symbiotic bacterium Mesorhizobium loti. DNA Res. 7:331– 338.

The Genus Brucella

443

Kannenberg, E. L., and R. W. Carlson. 2001. Lipid A and O-chain modifications causes Rhizobium lipopolysaccharides to become hydrophobic during bacteroid development. Molec. Microbiol. 39:379–391. Katz, S., E. Kowentz-Leutz, C. Mü ller, K. Meese, S. A. Ness, and A. Leutz. 1993. The NF-M transcription factor is related to C/EBP? and plays a role in signal transduction, differentiation and leukemogenesis of avian myelomonocytic cells. EMBO J. 12:1321–1332. Kaufmann, A. F., M. I. Meltzer, and G. P. Schmid. 1997. The economic impact of a bioterrorist attack: Are prevention and postattack intervention programs justifiable? Emerging Infect. Dis. 3:83–94. Keleti, G., D. S. Feingold, and J. S. Yongner. 1974. Interferon induction in mice by lipopolysaccharide from Brucella abortus. Infect. Immun. 10:182–283. Kelly, N. M., L. Young, and A. S. Cross. 1991. Differential induction of tumor necrosis factor expressing rough and smooth lipopolysaccharide phenotypes. Infect. Immun. 59:4491–4496. Kerkhofs, P., Y. Botton, P. Thiange, P. Dekeyser, and J. N. Limet. 1990. Diagnosis of bovine brucellosis by enzyme immunoassay of milk. Vet. Microbiol. 24:73–80. Kersh, G. J., E. N. Kersh, D. H. Fremont, and P. M. Allen. 1998. High and low-potency ligands with similar affinities for the TCR: The importance of kinetics in TCR signaling. Immunity 9:817–826. Kim, J., and J. E. Mayfield. 1997. Brucella abortus arginase and ornithine cyclodeaminase genes are similar to Ti plasmid arginase and ornithine cyclodeaminase. Biochim. Biophys. Acta 1354:55–57. Kim, J. A., Z. Sha, and J. E. Mayfield. 2000. Regulation of Brucella abortus catalase. Infect. Immun. 68:3681–3866. Kittelberger, R., F. Hilbink, M. F. Hansen, G. P. Ross, G. W. de Lisle, A. Cloeckaert, and J. de Bruyn. 1995a. Identification and characterization of immunodominant antigens during the course of infection with Brucella ovis. J. Vet. Diagn. Invest. 7:210–218. Kittelberger, R., F. Hilbink, M. F. Hansen, G. P. Ross, M. A. Joyce, S. Fenwick, J. Heesemann, H. Wolf-Watz, and K. H. Nielsen. 1995b. Serological cross-reactivity between Brucella abortus and Yersinia enterocolitica 0:9. II: The use of Yersinia outer proteins for the specific detection of Yersinia enterocolitica infections in ruminants. Vet. Microbiol. 47:271–280. Kittelberger, R., M. P. Reichel, M. A. Joyce, and C. Staak. 1997. Serological crossreactivity between Brucella abortus and Yersinia enterocolitica 0:9. III: Specificity of the in vitro antigen-specific gamma interferon test for bovine brucellosis diagnosis in experimentally Yersinia enterocolitica 0:9-infected cattle. Vet. Microbiol. 57:361– 371. Kittelberger, R., D. S. Diack, N. Vizcaino, M. S. Zygmunt, and A. Cloeckaert. 1998. Characterization of an immunodominant antigen in Brucella ovis and evaluation of its use in an enzyme-linked immunosorbent assay. Vet. Microbiol. 59:213–227. Knolle, P. A., T. Germann, U. Treichel, A. Uhrig, E. Schmitt, S. Hegenbarth, A. W. Lohse, G. Gerken. 1999. Endotoxin down-regulates T cell activation by antigenpresenting liver sinusoidal endothelial cells. J. Immunol. 162:1401–1407. Knosel, H., E. Hempel, and E. Forschner. 1991. Erfahrungen bei der Bekampfung der Brucellose in Rinderbestanden des Regierungsbezirks Hannover. DTW Deutsch. Tierarztl. Wochenschr. 98:356–358.

444

E. Moreno and I. Moriyó n

Köhler, S., J. Teyssier, A. Cloeckaert, B. Rouot, and J. P. Liautard. 1996. Participation of the molecular chaperone DnaK in intracellular growth of Brucella suis within U937-derived phagocytes. Molec. Microbiol. 20:701– 712. Kolst, A.-B. 1999. Time for a fresh look at the bacteria chromosome. Trends Microbiol. 7:223–226. Kolsto, A. B. 1997. Dynamic bacterial genome organization. Molec. Microbiol. 24:241–248. Kondo, S., K. Ishida, Y. Isshiki, Y. Haishima, T. Iguchi, and K. Hisatsune. 1993. N-3-hydroxypropionyl-alpha-Dperosamine homopolymer constituting the O-chain of lipopolysaccharides from Vibrio bioserogroup 1875 possessing antigenic factor(s) in common with O1 Vibrio cholerae. Biochem. J. 292:531–535. Kondo, S., Y. Sano, Y. Isshiki, and K. Hisatsune. 1996. The O polysaccharide chain of the lipopolysaccharide from Vibrio cholerae O76 is a homopolymer of N-[ (S)-(+)-2hydroxypropionyl]-alpha-L-perosamine. Microbiology 142:2879–2885. Kreutzer, D. L., and D. C. Robertson. 1979a. Surface macromolecules and virulence in intracellular parasitism: Comparison of cell envelope components of smooth and rough strains of Brucella abortus. Infect. Immun. 23:819–828. Kreutzer, D. L., L. A. Deyfus, and D. C. Robertson. 1979b. Interactions of polymorphonuclear leukocytes with smooth and rough strains of Brucella abortus. Infect. Immun. 23:737–742. Krieg, N. R., and J. G. Holt. 1984. Bergey’s Manual of Systematic Bacteriology. William and Wilkins. Baltimore, MD. 1: Krieger, J. I., S. F. Grammer, H. M. Grey, and R. W. Chesnut. 1985. Antigen presentation by splenic B cells: Resting B cells are ineffective, whereas activated B cells are effective accessory cells for T cell responses. J. Immunol. 135:2937–2945. Krueger, C. M., K. L. Marks, and G. M. Ihler. 1993. Bartonella bacilliformis genome size estimate and preliminary macrorestriction map. Abstracts of th 94th general meeting. Amer. Soc. Microbiol. H-187:233. Kü ndig, C., H. Hennecke, and M. Göttfert. 1994. Correlated physical and genetic map of the Bradyrhizobium japonicum 110 genome. J. Bacteriol. 175:613–622. Kuppuswamy, P. B. 1954. Chemoterapy of brucellosis in rams. NZ Vet. J. 2:110–113. Kurtz, R. S., and D. T. Berman. 1986. Influence of endotoxinprotein in immunoglobulin G isotype responses of mice to Brucella abortus lipopolysaccharide. Infect. Immun. 54:728–734. Kuzumawati, A., C. Cazevieille, F. Porte, S. Bettache, J. P. Liautard, and J. Sri-Widada. 2000. Early events and implication of F-actin and annexin I associated structures in the pathogenic uptake of Brucella suis by J774A1 murine cell line and human monocytes. Microb. Pathog. 28:343–352. Lacave, C., and J. Roux. 1965. Composition chimique des parois de Brucella: Glycosaminopeptide de Brucella abortus et Brucella melitensis. C. R. Acad. Sci. Paris 260:1514–1517. Laguerre, G., M. Bardin, and N. Amarger. 1993. Isolation from soil of symbiotic and nonsymbiotic Rhizobium leguminosarum by DNA hybridization. Can. J. Microbiol. 39:1142–1149. Laguerre, G., M. Bardin, and N. Amarger. 1994. Rapid identification of Rhizobia by restriction fragment length

CHAPTER 3.1.16 polymorphism analysis of PCR-amplified 16S rRNA genes. Appl. Environ. Microbiol. 60:56–62. Lambert, B., H. Joos, S. Dierickx, R. Vantomme, J. Swings, K. Kersters, and M. Van Montagu. 1990. Identification and plant interaction of Phyllobacterium sp., a predominant rhizobacterium of young sugar beet plants. Appl. Environ. Microbiol. 56:1093–1102. Lapham, C., B. Golding, J. Inman, R. Blackburn, J. Manischewitz, P. Highet, and H. Golding. 1996. Brucella abortus conjugated with a peptide derived from the V3 loop of human immunodeficiency virus (HIV) type 1 induces HIV-specific cytotoxic T-cell responses in normal and in CD4+ cell-depleted BALB/c mice. J. Virol. 70:3084–3092. Latimer, E., J. Simmers, N. Sriranganathan, R. M. Roop 2nd, G. G. Schurig, and S. M. Boyle. 1992. Brucella abortus deficient in copper/zinc superoxide dismutase is virulent in BALB/c mice. Microbiol. Pathogen. 12:105–113. Lebhun, M., W. Achouak, M. Scholter, O. Berge, H. Meier, M. Barakat, A. Hartmann, and T. Heulin. 2000. Taxonomic characterization of Ochrobactrum sp. isolates from soil samples and wheat roots and description of Ochrobactrum titrici sp. nov. and Ochrobactrum grigonense sp. nov. Int. J. Syst. Evol. Bacteriol. 50:2207– 2223. Lee, C. M., E. P. Mayer, J. Molnar, and M. Teodorescu. 1983. The mechanism of natural binding of bacteria to human lymphocyte subpopulations. J. Clin. Lab. Immunol. 11:87–94. Leiva-Leó n, J., M. de la Rosa, J. Plata, M. J. Martínez, and A. J. Jiménez. 1991. An immunoblotting study of serologic response in patients with acute brucellosis. Diagn. Microbiol. Infect. Dis. 14:515–518. Leonard, B. A., I. Ló pez-Goñ i, and C. L. Baldwin. 1997. Brucella abortus siderophore 2,3-dihidroxybenzoic acid protects brucellae from killing by macrophages. Vet. Res. 28:87–92. Letesson, J. J., A. Tibor, G. van Eynde, V. Wansard, V. E. Weynants, P. A. Denoel, and E. Saman. 1997. Humoral immune responses of Brucella-infected cattle, sheep, and goats to eight purified recombinant Brucella proteins in an indirect enzyme-linked immunosorbent assay. Clin. Diagn. Lab. Immunol. 4:556–564. LeVier, K., R. W. Phillips, V. K. Grippe, R. M. Roop, and G. C. Walker. 2000. Similar requirements of a plant symbiont and a mammalian pathogen for prolonged intracellular survival. Science 287:2492–2493. Levin, B. R., and R. E. Lenski. 1983. Coevolution in bacteria, their viruses and plasmids. In: D. J. Futiyama and M. Statkin (Eds.) Coevolution. Sinauer Associates Publishers. Suderland, MA. 99–127. Liautard, J. P., A. Gross, J. Dornand, and S. Köhler. 1996. Interactions between professional phagocytes and Brucella spp. Microbiologia 12:197–206. Lin, J., L. G. Adams, and T. A. Ficht. 1992. Characterization of the heat shock response in Brucella abortus and isolation of the genes encoding the GroE heat shock proteins. Infect. Immun. 60:2425–2431. Lin, J., and T. A. Ficht. 1995. Protein synthesis in Brucella abortus induced during macrophage infection. Infect. Immun. 63:1409–1414. Lin, J., L. G. Adams, and T. A. Ficht. 1996. Immunological response of the Brucella abortus GroEL homologue. Infect. Immun. 64:4396–4400. Lindberg, A. A., S. Haeggman, K. Karlson, H. E. Carlsson, and N. S. Mair. 1982. Enzyme immunoassay of the anti-

CHAPTER 3.1.16 body response to Brucella and Yersinia enterocolitica 09 infections in humans. J. Hyg. (Lond.) 88:295–307. Liou, W., H. J. Geuze, M. J. H. Geelen, and J. W. Slot. 1997. The autophagic and endocytic pathways converge at the nascent autophagic vacuoles. J. Cell Biol. 136:61–70. Long, S. R. 1989. Rhizobium genetics. Ann. Rev. Genet. 23:483–506. Lopetegui P. 1999. Consulta de expertos en vacunació n frente a brucelosis y estrategias de vacunació n de brucelosis en programas de control y erradicació n PAHO/ WHO publications. Chile. 16–18. Ló pez-Goñ i, I., I. Moriyó n, and J. B. Neilands. 1992. Identification of 2,3. dihidroxybenzoic acid as Brucella abortus siderophore. Infect. Immun. 60:4496–4503. n, A. Ló pez-Urrutia, L., A. Alonso, M. L. Nieto, Y. Bayó Orduñ a, and M. S. Crespo. 2000. Lipopolysaccharide of Brucella abortus and Brucella melitensis induce oxide synthesis in rat peritoneal macrophages. Infect. Immun. 68:1740–1745. Lord, V. R., M. R. Rolo, and J. W. Cherwonogrodzky. 1989. Evaluation of humoral immunity to Brucella sp in cattle by use of an agar-gel immunodiffusion test containing a polysaccharide antigen. Amer. J. Vet. Res. 50:1813–1816. Lord, V. R., J. W. Cherwonogrodzky, G. G. Schurig, R. D. Lord, M. J. Marcano, and G. E. Melendez. 1998a. Venezuelan field trials of vaccines against brucellosis in swine. Amer. J. Vet. Res. 59:546–551. Lord, V. R., G. G. Schurig, J. W. Cherwonogrodzky, M. J. Marcano, and G. E. Melendez. 1998b. Field study of vaccination of cattle with Brucella abortus strains RB51 and 19 under high and low disease prevalence. Amer. J. Vet. Res. 59:1016–1020. Lowenstein, C. J., E. W. Alley, P. Raval, A. M. Snowman, S. H. Snyder, S. W. Russell, and W. J. Murphy. 1993. Macrophage nitric oxide synthase gene: Two upstream regions mediate induction by interferon gamma and lipopolysaccharide. Proc. Natl. Acad. Sci. USA 90:9730– 9734. Lucero, N. E., L. Foglia, S. M. Ayala, D. Gall, and K. Nielsen. 1999. Competitive enzyme immunoassay for diagnosis of human brucellosis. J. Clin. Microbiol. 37:3245–3248. Mackaness, G. B. 1967. The relationship of delayed hypersensitivity to acquired cellular resistance. Br. Med. Bull. 23:52–54. Mackenzie, C., M. Chidambaram, E. J. Sodergren, S. Kaplan, and G. M. Weinstock. 1995. DNA repair mutants of Rhodobacter sphaeroides. J. Bacteriol. 177:3027–3035. Macmillan, A. P. 1990a. Conventional serological tests. In: K. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 153–197. Macmillan, A. P., I. Greiser-Wilke, V. Moennig, and L. A. Mathias. 1990b. A competition enzyme immunoassay for brucellosis diagnosis. DTW Deutsch. Tierarztl. Wochenschr. 97:83–85. Maeno N. H. Oda, K. Yoshiie, M. R. Wahid, T. Fujimura, and S. Matayoshi. 1999. Live Bartonella henselae enhances endothelial cell proliferation without direct contact. Microb. Pathog. 27:419–427. Maeno, N., K. Yoshiie, S. Matayoshi T. Fujimura, S. Mao, M. R. Wahid, and H. Oda. 2002. A heat-stable component of Bartonella adhesion molecule-1 expression on vascular endothelial cells. Scan. J. Immunol. 55:366–372. Manthei, C. A., D. E. DeTray, and E. R. Goode. 1950. Brucella infection in bulls and spread of brucellosis in cattle by artificial insemination. Proc. Amer. Vet. Med. Assoc. 87:177.

The Genus Brucella

445

Mardarowicz, C. 1966. Isolierung und Charakterisierung des Murein-sacculus von Brucella. Z. Naturforschg. 21b:1006–1007. Marín, C. M., M. P. Jiménez de Bagü és, M. Barberán, and J. M. Blasco. 1989a. Efficacy of long-acting oxytetracycline alone or in combination with streptomycin for treatment of Brucella ovis infection of rams. Amer. J. Vet. Res. 50:560–563. Marín, C. M., M. P. Jiménez de Bagü és, J. M. Blasco, C. Gamazo, I. Moriyó n, and R. Díaz. 1989b. Comparison of three serological tests for Brucella ovis infection of rams using different antigenic extracts. Vet. Rec. 125:504–508. Marín, C. M., M. Braberán, M. P. Jiménez de Bagü és, and J. M. Blasco. 1990. Comparison of subcutaneous and conjunctival routes of Rev 1 vaccination for the prophylaxis of Brucella ovis infection in rams. Res. Vet. Sci. 48:209–215. Marín, C. M., M. P. Jiménez de Bagü és, M. Braberán, and J. M. Blasco. 1996. Comparison of two selective media for the isolation of Brucella melitensis from naturally infected sheep and goats. Vet. Rec. 138:409–411. Marín, C. M., B. Alonso-Urmeneta, I. Moriyó n, S. PérezGó mez, and J. M. Blasco. 1998. Comparison of polyclonal, monoclonal and protein G peroxidase conjugates in an enzyme-linked immunosorbent assay for the diagnosis of Brucella ovis in sheep. Vet. Rec. 143:390– 394. Marín, C. M., E. Moreno, I. Moriyó n, R. Díaz, and J. M. Blasco. 1999. Performance of competitive and indirect enzyme-linked immunosorbent assay, gel immunoprecipitation with native hapten polysaccharide and standard serological tests in the diagnosis of sheep brucellosis. Clin. Diagn. Lab. Immunol. 6:269–272. Marquis, H., and T. A. Ficht. 1993. The omp2 gene locus of Brucella abortus encodes two homologous membrane proteins with properties characteristic of porins. Infect. Immun. 61:3785–3790. Marr, A. G., and J. B. Wilson. 1950. Characteristics of carbondioxide-independent cultures of Brucella abortus isolated from cattle vaccinated with strain 19 (cited by Jones, L. M., V. Montgomery, and J. B. Wilson). J. Infect. Dis. 115:312–320. Marr, A. G., C. B. Olsen, C. B. Unger, and J. B. Wilson. 1953. The oxidation of glutamic acid by Brucella abortus. J. Bacteriol. 66:606–610. Martínez de Tejada, G., and I. Moriyó n. 1993. The outer membranes of Brucella spp. are not barriers to hydrophobic permeants. J. Bacteriol. 175:5273–5275. Martínez de Tejada, G., J. Pizarro, E. Moreno, and I. Moriyó n. 1995. The outer membranes of Brucella spp. are resistant to bactericidal cationic peptides. Infect. Immun. 63:3054–3061. Maruyama S. R., W. Kasten, H. J. Boulouis, N. A. Gurfield, Y. Katsube, and B. B. Chomel. 2001. Genomic diversity of Bartonella henselae isolates from domestic cats from Japan, the USA and France by pulsed-field gel electrophoresis. Vet. Microbiol. 79:337–349. Masterson, R. V., R. K. Prakash, and A. G. Atherly. 1985. Conservation of symbiotic nitrogen fixation sequences in Rhizobium japonicum and Bradyrhizobium japonicum. J. Bacteriol. 163:21–26. Matar, G. M., I. A. Khneisser, and A. M. Abdelnoor. 1996. Rapid laboratory confirmation of human brucellosis by PCR analysis of a target sequence on the 31-kilodalton Brucella antigen DNA. J. Clin. Microbiol. 34:477–478.

446

E. Moreno and I. Moriyó n

Maurin, M., V. Roux, A. Stein, F. Ferrier, R. Viraben, and D. Raoult. 1994. Isolation and characterization by immunofluorescence, sodium dodecyl sulfate-polyacrylamide gel electrophoresis, western blot, restriction fragment length polymorphism-PCR, 16SrRNA gene sequencing, and pulse gel electrophoresis of Rochalimaea quintana from a patient with bacillary angiomatosis. J. Clin. Microbiol. 32:1166–1171. Mayfield, J. E., J. A. Bantle, D. R. Ewalt, V. P. Meador, and L. B. Tabatabai. 1990. Detection of Brucella cells and cell components. In: K. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 98– 120. McCorquodale, S. M., and R. F. DiGiacomo. 1985. The role of wild North American ungulates in the epidemiology of bovine brucellosis: A review. J. Wildl. Dis. 21:351–357. McCullough, N. B., and L. A. Dick. 1943. Growth of Brucella in a simple chemically defined medium. Proc. Soc. Exp. Biol. Med. 52:310–311. McCullough, N. B., and G. A. Beal. 1951. Growth and manometric studies on carbohydrate utilization of Brucella. J. Infect. Dis. 89:266–271. McQuiston, J. R., R. Vemulapalli, T. J. Inzana, G. G. Schurig, N. Sriranganathan, D. Fritzinger, T. L. Hadfield, R. A. Warren, N. Snellings, D. Hoover, S. M. Halling, and S. M. Boyle. 1999. Genetic characterization of a Tn5disrupted glycosyltransferase gene homolog in Brucella abortus and its effect on lipopolysaccharide composition and virulence. Infect. Immun. 67:3830–3835. Mercier, E., B. E. Jumas, S. A. Allardet, D. O’Callaghan, and M. Ramuz. 1996. Polymorphism in Brucella strains detected by studying distribution of two short repetitive DNA elements. J. Clin. Microbiol 34:1299–1302. Méresse, S., O. Steele-Mortimer, E. Moreno, M. Desjardins, B. Finlay, and J. P. Gorvel. 1999. Controlling the maturation of pathogen-containing vacuoles: Matter of life and death. Nature Cell Biol. 1:E183–E188. Merrel, B. R., E. Weiss, and G. A. Dasch. 1978. Morphological and cell association characteristics of Rochalimea quintana: Comparison of the Vole and Fuller strains. J. Bacteriol. 135:633–640. Meyer, M. E. 1976. Evolution and taxonomy in the genus Brucella: Steroid hormone induction of filterable forms with altered characteristics after reversion. Amer. J. Vet. Res. 37:207–210. Meyer, M. E. 1977. Epidemiological odds and ends. In: R. P. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 135– 142. Meyer, M. E. 1985. Characterization of Brucella abortus strain 19 isolated from human and bovine tissues and fluids. Amer. J. Vet. Res. 46:902–904. Meyer, M. E. 1990. Current concepts in the taxonomy of the genus Brucella. In: K. Nielsen, and B. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 1–27. Michaux-Charachon, S., J. Paillisson, M. G. Carles-Nurit, G. Bourg, A. Allardet-Servent, and M. Ramuz. 1993. Presence of two independent chromosomes in the Brucella melitensis 16M genome. J. Bacteriol. 175:701–705. Michaux-Charachon, S., G. Bourg, E. Jumas-Bilak, P. Guigue-Talet, A. Allardet-Servent, D. O’Callaghan, and M. Ramuz. 1997. Genome structure and phylogeny in the genus Brucella. J. Bacteriol. 179:3244–3249. Mikolon, A. B., I. A. Gardner, S. K. Hietala, J. Hernández de Anda, E. Chamizo-Pestana, S. G. Hennager, and A. J.

CHAPTER 3.1.16 Edmondson. 1998. Evaluation of North American antibody detection tests for diagnosis of brucellosis in goats. J. Clin. Microbiol. 36:1716–1722. Milward, F. W., P. Nicoletti, and E. Hoffmann. 1984. Effectiveness of various therapeutic regimens for bovine brucellosis. Amer. J. Vet. Res. 45:1825–1828. Mittal, K. R., and I. R. Tizard. 1979. A simple technique to differentiate between animals infected with Yersinia enterocolitica IX and those infected with Brucella abortus. Res. Vet. Sci. 26:248–250. Mittal, K. R., and I. R. Tizard. 1980a. Studies on the relationship between Yersinia enterocolitica IX and Brucella abortus agglutinins in naturally infected animals. Res. Vet. Sci. 28:311–314. Mittal, K. R., I. D. Ricciardi, and I. R. Tizard. 1980b. Indirect hemagglutination employing enterobacterial common antigen and Yersinia somatic antigen: A technique to differentiate brucellosis from infections involving cross-reacting Yersinia enterocolitica. J. Clin. Microbiol. 11:149–152. Mittal, K. R., I. R. Tizard, and D. A. Barnum. 1985. Serological cross-reactions between Brucella abortus and Yersinia enterocolitica 0:9. Int. J. Zoonoses 12:219– 227. Moore, C. G., and P. R. Schnurrenberger. 1981. A review of naturally ocurring Brucella abortus infections in wild mammals. J. Amer. Vet. Assoc. 179:779–782. Morata, P., M. I. Queipo-Ortuñ o, and J. D. Colmenero. 1998. Strategy for optimizing DNA amplification in a peripheral blood PCR assay used for diagnosis of human brucellosis. J. Clin. Microbiol. 36:2443–2446. Morata, P., M. I. Queipo-Ortuñ o, and J. D. Colmenero. 1999a. Author’s reply. J. Clin. Microbiol. 37:1654–1655. o, J. M. Reguera, O. M. Morata, P., M. I. Queipo-Ortuñ Garcia, C. Pichardo, and J. D. Colmenero. 1999b. Posttreatment follow-up of brucellosis by PCR assay. J. Clin. Microbiol. 37:4163–4166. Moreno, E., and D. T. Berman. 1979. Brucella abortus lipopolysaccharide is mitogenic for spleen cells od endotoxin-resistant C3H/HeJ mice. J. Immunol. 123:2915–2919. Moreno, E., D. T. Berman, and L. A. Boettcher. 1981. Biological activities of Brucella abortus lipopolysaccharides. Infect. Immun. 31:362–370. Moreno, E., L. M. Jones, and D. T. Berman. 1984a. Immunochemical characterization of rough Brucella lipopolysaccharides. Infect. Immun. 43:779–782. Moreno, E., R. S. Kurtz, and D. T. Berman. 1984b. Induction of immune and adjuvant immunoglobulin G responses in mice by Brucella lipopolysaccharide. Infect. Immun. 46:74–80. Moreno, E., H. Mayer, and I. Moriyó n. 1987. Characterization of a native polysaccharide hapten from Brucella melitensis. Infect. Immun. 55:2850–2853. Moreno, E., E. Stackebrandt, M. Dorsch, J. Wolters, M. Busch, and H. Mayer. 1990. Brucella abortus 16S rRNA and lipid A reveal a phylogenetic relationship with members of the alpha-2 subdivision of the class Proteobacteria. J. Bacteriol. 172:3569–3576. Moreno, E. 1992. Brucella evolution in Prevention of Brucellosis in Mediterranean Countries. In: M. Plommet (Ed.) International Centre for Advanced Mediterranean Agronomic Studies/Pudoc Scientific Publishers. Wageningen, The Netherlands. 198–218. Moreno, E. 1997. In search of bacterial species definition. Rev. Biol. Trop. 45:753–771.

CHAPTER 3.1.16 Moreno, E. 1998. Genome evolution within the alpha Proteobacteria: Why do some bacteria not possess plasmids and others exhibit more than one different chromosome? FEMS Microbiol. Rev. 22:255–275. Moreno, E., and I. Moriyó n. 2002. Brucella melitensis: A nasty bug with hidden credentials for virulence. Proc. Nat. Acd. Sci. USA 99 :1–3. Morgan, W. J. B. 1977. The diagnosis of Brucella abortus infections in Britain. In: R. P. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 21–39. Moriyó n, I., and D. T. Berman. 1982. Effects of nonionic, ionic, and dipolar ionic detergents and EDTA on the Brucella cell envelope. J. Bacteriol. 152:822–828. Moriyó n, I., C. Gamazo, and R. Díaz. 1987. Properties of the outer membrane of Brucella. Ann. Inst. Pasteur Microbiol. 138:89–91. Moriyó n, I., and I. Ló pez-Goñ i. 1998. Structure and properties of the outer membranes of Brucella abortus and Brucella melitensis. Int. Microbiol. 1:19–26. Moulton, J. E., and M. E. Meyer. 1958. The pathogenesis of Brucella suis infection in guinea pigs. Lesions of the spleen, liver, testis, and articulations. Cornell. Vet. 48:165–172. Moyer, N. P., and W. J. Hausler Jr. 1992. The genus Brucella. In: A. Balows, H. G. Trü per, M. Dworkin, W. Harder and K. H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York, NY. 2384–2400. Mustafa, A. A., and M. Abusowa. 1993. Field-oriented trial of the Chinese Brucella suis strain 2 vaccine on sheep and goats in Libya. Vet. Res. 24:422–429. Myers, D. M., and A. Siniuk. 1970. Preliminary report on the development of a diffusion in gel methods for the diagnosis of ram epididimitis. Appl. Microbiol. 19:335– 337. Naroeni, A., N. Jouy, S. Ouahrani-Bettache, J. P. Liautard, and F. Porte. 2001. Brucella suis-impaired specific recognition of phagosome by lysosomes due to phagosomal membrane modifications. Infect. Immun. 69:486– 493. Naroeni, A., and F. Porte. 2002. Role of cholesterol and the ganglioside GM(1) in entry and short-term survival of Brucella suis in murine macrophages. Infect. Immun. 70:1640–1644. Navarro, E., J. A. Fernández, J. Escribano, and J. Solera. 1999. PCR assay for diagnosis of human brucellosis. J. Clin. Microbiol. 37:1654–1655. Ne’eman, L., and L. M. Jones. 1963. The evaluation of the virulence of a recently isolated strain of Brucella melitensis (Y 85) for guinea pigs and pregnant sheep. Refu. Vet. 20:169–172. Neumeister, B., M. Faigle, M. Sommer, U. Zähringer, F. Stelter, R. Menzel., S. Schü tt, and H. Northoff. 1988. Low endotoxic potential of Legionella pneumophila lipopolysaccharide due to failure of interaction with the monocyte lipopolysaccharide receptor CD14. Infect. Immun. 66:4151–4157. Newton, J. W., and J. B. Wilson. 1954. Carbon dioxide requirements and nucleic acid synthesis in Brucella abortus. J. Bacteriol. 68:74–76. Newton, J. W., A. G. Marr, and J. B. Wilson. 1954b. Fixation of carbon14 dioxide into nucleic acid constituents by Brucella abortus. J. Bacteriol. 67:236. Nicoletti, P. 1977. Adult vaccination. In: R. P. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 201–208.

The Genus Brucella

447

Nicoletti, P. L., L. M. Jones, and D. T. Berman. 1978. Comparison of the subcutaneous and conjunctival route of vaccination with Brucella abortus strain 19 vaccine in adult cattle. J. Am. Vet. Med. Assoc. 173:1450–1456. Nicoletti, P., F. W. Milward, E. Hoffmann, and L. Altvater. 1985. Efficacy of long-acting oxytetracycline alone or combined with streptomycin in the treatment of bovine brucellosis. J. Am. Vet. Med. Assoc. 187:493–495. Nicoletti, P., R. P. Lenk, M. C. Popescu, and C. E. Swenson. 1989. Efficacy of various treatment regimens, using liposomal streptomycin in cows with brucellosis. Amer. J. Vet. Res. 50:1004–1007. Nicoletti, P. L. 1990a. Vaccination. In: K. H. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 283–299. Nicoletti, P., and A. J. Winter. 1990. The immune response to Brucella abortus— the cell-mediated response to infections: Antigens of Brucella. In: K. Nielsen and B. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 83–95. Nielsen, K., and J. R. Duncan. 1988. Antibody isotype response in adult cattle vaccinated with Brucella abortus strain 19. Vet. Immunol. Immunopathol. 19:205–209. Nielsen, K. H. 1990a. The serological response of cattle immunized with Yersinia enterocolitica O:9 or O:16 to Yersinia and Brucella abortus antigens in enzyme immunoassays. Vet. Immunol. Immunopathol. 24:373–382. Nielsen, K., and B. Duncan. 1990b. Animal Brucellosis. CRC Press. Boca Raton, FL. Nielsen, K., L. Kelly, D. E. Gall, P. L. Nicoletti, and W. Kelly. 1995. Improved competitive enzyme immunoassay for the diagnosis of bovine brucellosis. Vet. Immunol. Immunopathol. 46:285–291. Nielsen, K., D. Gall, M. Jolley, G. Leishman, S. Balsevicius, P. Smith, P. Nicoletti, and F. Thomas. 1996a. A homogeneous fluorescence polarization assay for detection of antibody to Brucella abortus. J. Immunol. Meth. 195:161–168. Nielsen, K. H., L. Kelly, D. Gall, S. Balsevicius, J. Bosse, P. L. Nicoletti, and W. Kelly. 1996b. Comparison of enzyme immunoassays for the diagnosis of bovine brucellosis. Prev. Vet. Med. 26:7–32. Nielsen, K., D. Gall, P. Smith, A. M. Vigliocco, B. Perez, L. Samartino, P. Nicoletti, A. Dajer, P. Elzer, and F. Enright. 1999. Validation of the fluorescence polarization assay as a serological test for the presumptive diagnosis of porcine brucellosis. Vet. Microbiol. 68:245– 253. Nierman W. C., T. V. Feldblyum, M. T. Laub, I. T. Paulsen, K. E. Nelson, J. A. Eisen, J. F. Heidelberg, M. R. Alley, N. Ohta, J. R. Maddock, I. Potocka, W. C. Nelson, A. Newton, C. Stephens, N. D. Phadke, B. Ely, R. T. DeBoy, R. J. Dodson, A. S. Durkin, M. L. Gwinn, D. H. Haft, J. F. Kolonay, J. Smit, M. B. Craven, H. Khouri, J. Shetty, K. Berry, T. Utterback, K. Tran, A. Wolf, J. Vamathevan, M. Ermolaeva, O. White, S. L. Salzberg, J. C. Venter, L. Shapiro, and C. M. Fraser. 2001. Complete genome sequence of Caulobacter crescentus. Proc. Natl. Acad. Sci. USA 98 :4136–4141. Nikolskaya, T., M. Fonstein, and R. Haselkorn. 1995. Alignment of a 1,2 Mb chromosomal region from three strains of Rhodobacter capsulatus reveals a significantly mosaic structure. Proc. Natl. Acad. Sci. USA 92:10609–10613. Oberti, J., R. Caravano, and J. Roux. 1982. Demonstration of an external layer in several species of the genus Brucella. Can. J. Microbiol. 28:1300–1303.

448

E. Moreno and I. Moriyó n

O’Callaghan, D., C. Cazevielle, A. Allardet-Servent, M. L. Boschiroll, G. Bourg, V. Foulongne, P. Frutos, Y. Kulakov, and M. Ramuz. 1999. A homologue of Agrobacterium tumefaciens VirB and Bordetella pertussis Ptl type IV systems is essential for intracellular survival of Brucella suis. Molec. Microbiol. 33:1210–1220. Ochman, H., and A. Wilson. 1987. Evolution in bacteria: Evidence for a universal substitution rate in Cellular Genomes. J. Molec. Evol. 26:74–86. O’Hara, R. J. 1994. Evolutionary history and the species problem. Amer. Zoologist 34:12–22. Oliveira, S. C., J. S. Harms,. E. L. Rech, R. S. Rodarte, A. L. Bocca, A. M. Goes, and G. A. Splitter. 1998. The role of T cell subsets and cytokines in the regulation of intracellular bacterial infection. Brazil. J. Med. Res. 31:77–84. Olsen, S. C. 2000. Immune responses and efficacy after administration of a commercial Brucella abortus strain RB51 vaccine to cattle. Vet. Therap. 1:183–191. Orci, L., B. S. Glick, and J. E. Rothman. 1986. A new type of coated vesicular carrier that appears not to contain clathrin: Its possible role in protein transport within the Golgi stack. Cell 46:171–184. Orci, L., M. Stamnes, M. Ravazzola, M. Amherdt, A. Perrelet, T. H. Sollner, and J. E. Rothman. 1997. Bidirectional transport by distinct populations of COPIcoated vesicles. Cell 90:335–349. Otero, J. R., A. Fuertes, E. Palenque, and A. R. Noriega. 1982. Microtiter-adapted method that facilitates the Coombs test for brucellosis. J. Clin. Microbiol. 16:737– 738. Ottones, F., J. Dornand, A. Naroeini, J.-P. Liautard, and J. Favero. 2000a. V gamma 9V delta 2 T cells impair intracellular multiplication of Brucella suis in autologous monocytes through soluble factor release and contact-dependent cytotoxic effect. J. Immunol. 165:7133–7239. Ottones, F., J. Liautard, A. Gross, F. Rabenoelina, J. P. Liautard, and J. Favero. 2000b. Activation of human V gamma9delta2 T cells by a Brucella suis non-petide fraction impairs bacterial intracellular multiplication in monocytic infected cells. Immunology 100:252–258. Ouahrani, S., S. Michaux, J. S. Widada, G. Bourg, R. Tournebize, M. Ramuz, and J. P. Liautard. 1993. Identification and sequence analysis of IS6501, an insertion sequence in Brucella spp. Relationship between genomic structure and number if IS6501 copies. J. Gen. Microbiol. 139:3265–3273. Ouahrani, S., M. P. Soubrier, and J. P. Liautard. 1996. IS6501anchored PCR for the detection and identification of Brucella species and strains. J. Appl. Bacteriol. 81:154– 160. Palmer, M. V., N. F. Cheville, and A. E. Jensen. 1996. Experimental infection of pregnant cattle with the vaccine candidate Brucella abortus strain RB51: pathologic, bacteriologic, and serologic findings. Vet. Pathol. 33:682– 691. Palmer, M. V., S. C. Olsen, and N. F. Cheville. 1997. Safety and immunogenicity of Brucella abortus strain RB51 vaccine in pregnant cattle. Amer. J. Vet. Res. 58:472– 477. Paquet, J. Y., M. A. Diaz, S. Genevrois, M. Grayon, J. M. Verger, X. De-Bolle, J. H. Lakey, J. J. Letesson, and A. Cloeckaert. 2001. Molecular, antigenic, and functional analyses of Omp2b porin size variants of Brucellas. J. Bacteriol. 183:4839–4847.

CHAPTER 3.1.16 Páramo, L., B. Lomonte, J. Pizarro-Cerdá, J. A. Bengoechea, J. P. Gorvel, and E. Moreno. 1998. Bactericidal activity of Lys49 and Asp49 myotoxic phospholipases A2 from Bothrops asper snake venom: Synthetic Lys49 myotoxin II-(115-129)-peptide identifies its bactericidal region. Eur. J. Biochem. 253:452–461. Paulsen I. T., R. Seshadri, K. E. Nelson, J. A. Eisen, J. F. Heidelberg, T. D. Read, R. J. Dodson, L. Umayam, L. M. Brinkac, M. J. Beanan, S. C. Daugherty, R. T. Deboy, A. S. Durkin, J. F. Kolonay, R. Madupu, W. C. Nelson, B. Ayodeji, M. Kraul, J. Shetty, J. Malek, S. E. Van Aken, S. Riedmuller, H. Tettelin, S. R. Gill, O. White, S. L. Salzberg, D. L. Hoover, L. E. Lindler, S. M. Halling, S. M. Boyle, C. M. Fraser. 2002. The Brucella suis genome reveals fundamental similarities between animal and plant pathogens and symbionts. Proc. Natl. Acad. Sci. USA 99:13148–13153. Pellicer, T., J. Ariza, A. Foz, R. Pallares, and F. Gudiol. 1988. Specific antibodies detected during relapse of human brucellosis. J. Infect. Dis. 157:918–924. Perry, M. B., and D. R. Bundle. 1990a. Antigenic relationships of the lipopolysaccharides of Escherichia hermannii strains with those of Escherichia coli O157:H7, Brucella melitensis, and Brucella abortus. Infect. Immun. 58:1391–1395. Perry, M. B., and D. R. Bundle. 1990b. Lipopolysaccharide antigens and carbohydrates of Brucella. In: L. G. Adams (Eds.) Advances in Brucellosis Research. Texas A&M University Press. College Station, TX. 76–88. Phillips, R. W., P. H. Elzer, and R. M. Roop 2nd. 1995. A Brucella melitensis high temperature requirement A (HtrA) deletion mutant demonstrates a stress response defective phenotype in vitro and transient attenuation in the BALB/c mouse model. Microb. Pathog. 19:227– 234. Phillips, R. W., P. H. Elzer, G. T. Robertson, S. D. Hagious, J. V. Walker, M. B. Fatemi, F. M. Enright, and R. M. Roop 2nd. 1997. A Brucella melitensis high-temperaturerequirement A (HtrA) deletion mutant is attenuated in goats and protects against abortion. Res. Vet. 63:165– 167. Pietz, D. E. 1977. Brucella antigens and serological tests results. In: R. M. Crawford and R. J. Hidalgo (Eds) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 49–60. Pinochet, L., P. Abalos, M. L. Sánchez, I. Palavicino, and M. A. Vent. 1989. Preparació n y evaluació n de un antígeno para descartar respuesta post-vacunal a Brucella abortus cepa 19. Av. Cs. Vet. 4:43–48. Pitt, A., L. S. Mayorga, P. L. Stahl, and A. L. Schwartz. 1992. Alterations in the protein composition of maturing phagosomes. J. Clin. Invest. 90:1978–1983. Pitt, T. L. 1994. Bacterial typing: The way ahead. J. Med. Microbiol. 40:1–2. Pizarro-Cerdá, J., M. Desjardins, E. Moreno, and J. P. Gorvel. 1997. When intracellular pathogens invade the frontiers of cell biology and immunology. Histol. Histopathol. 12:1027–1038. Pizarro-Cerdá, J. 1998a. Traffic intracellulaire ET survie de Brucella abortus dans les phagocytes professionnels et non professionnels (doctoral thesis). Université de la Mediterranée Aix-Marseille II, Faculté des Sciences de Luminy. Marseille-Luminy , France. 156–168. Pizarro-Cerdá, J., S. Meresse, R. G. Parton, G. Van der Goot, A. Sola-Landa, I. Ló pez-Goñ i, E. Moreno, and J. P. Gorvel. 1998b. Brucella abortus transits through the

CHAPTER 3.1.16 autophagic pathway and replicates in the endoplasmic reticulum of nonprofessional phagocytes. Infect. Immun. 66:5711–5724. Pizarro-Cerdá, J., E. Moreno, V. Sanguedolce, J. L. Mege, and J. P. Gorvel. 1998c. Virulent Brucella abortus prevents lysosome fusion and is distributed within autophagosome-like compartments. Infect. Immun. 66:2387– 2392. Pizarro-Cerdá, J., M. Desjardins, E. Moreno, S. Akira, and J. P. Gorvel. 1999a. Modulation of endocytosis in nuclear factor IL-6 (-/-) macrophages is responsible for a high susceptibility to intracellular bacterial infection. J. Immunol. 162:3519–3526. Pizarro-Cerdá, J., E. Moreno, and J. P. Gorvel. 1999b. Brucella abortus invasion and survival within professional and nonprofessional phagocytes. Microb. Infect. 6:201– 232. Pizarro-Cerdá, J., E. Moreno, and J. P. Gorvel. 2000. Invasion and intracellular trafficking of Brucella abortus in nonphagocytic cells. Adv. Cell. Molec. Biol. Membr. Organelles 2:829–835. Plenderleith, R. W. 1970. Some observations on brucellosis in a Jersey herd, 1965–1969. Vet. Rec. 87:40–46. Plommet, M. 1977. Studies of experimental brucellosis in cows in France. In: R. M. Crawford and R. J. Hidalgo (Eds.) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 116–1134. Plommet, M., and R. Fensterbank. 1984. La vaccination antibrucellique administrée par voie conjonctivale. Dev. Biol. Stand. 56:681–687. Plommet, M. 1991. Minimal requirements for growth of Brucella suis and other Brucella species. Zentralbl. Bakteriol. 275:436–450. Plommet, M. 1992. Prevention of Brucellosis in Mediterranean Countries. International Centre for Advanced Mediterranean Agronomic Studies/Pudoc Scientific Publishers. Wageningen , The Netherlands. 1–289. Poindexter, J. S. 1989. Genus Caulobacter, p. 1924–1939. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. G. Holt (ed.). Bergey’s manual of Systematic Bacteriology Williams & Wilkins. Baltimore, MD. Pomales-Lebron, A., and W. R. Stinebring. 1957. Intracellular multiplication of Brucella abortus in normal and immune mononuclear phagocytes. Proc. Soc. Exp. Biol. Med. 94:78–83. Poquet, Y., M. Kroca, F. Halary, S. Stenmark, M.-A. Peyrat, M. Bonneville, J.-J. Fournié, and A. Sjöstedt. 1998. Expansion of Vγ9Vδ2 T cells is triggered by Francisella tularensis-derived phosphoantigens in tularemia vaccine. Infect. Immun. 66:2107–2112. Porte, F., J. P. Liautard, and S. Kohler. 1999. Early acidification of phagosomes containing Brucella suis is essential for intracellular survival in murine macrophages. Infect. Immun. 67:4041–4047. Prakosh, R. K., and R. A. Schilperoort. 1982. The relationship between nif plasmids of fast growing Rhizobium species and Ti plasmids of Agrobacterium tumefaciens. J. Bacteriol. 149:1129–1134. Price, R. E., J. W. Templeton, and L. G. Adams. 1990. Survival of smooth, rough and transposon mutant strains of Brucella abortus in bovine mammary macrophages. Vet. Immunol. Immunopathol. 26:353–365. Queipo-Ortuñ o, M. I., P. Morata, P. Ocó n, P. Manchado, and J. D. Colmenero. 1997. Rapid diagnosis of human brucellosis by peripheral-blood PCR assay. J. Clin. Microbiol. 35:2927–2930.

The Genus Brucella

449

Queipo-Ortuñ o, M. I., M. A. García-Ordó ez, J. D. Colmenñ ero, and P. Morata. 1999. Hydrogen peroxide improves the efficiency of a peripheral blood PCR assay for diagnosis of human brucellosis. Biotechniques 27:248– 252. Rabinowitz, J. D., C. Beeson, D. S. Lyons, M. M. Davis, and H. M. McConnell. 1996. Kinetic discrimination in T-cell activation. Proc. Natl. Acad. Sci. USA 93:1401–1405. Radwan, A. I., S. I. Bekairi, A. M. al-Bokmy, P. V. Prasad, O. M. Mohamed, and S. T. Hussain. 1993. Successful therapeutic regimens for treating Brucella melitensis and Brucella abortus infections in cows. Rev. Sci. Tech. 12:909–922. Rafie-Kolpin, M., R. C. Essenberg, and J. H. Wyckoff. 1996. Identification and comparison of macrophage-induced proteins and proteins induced under various stress conditions in Brucella abortus. Infect. Immun. 64:5274– 5283. Rasool, O., E. Freer, E. Moreno, and C. Jarstrand. 1992. Effect of Brucella abortus lipopolysaccharide on oxidative metabolism and lysozyme release by human neutrophils. Infect. Immun. 60:1699–1702. Rava, P. S., L. Somma, and H. M. Steinman. 1999. Identification of a regulator that controls stationary-phase expression of catalase-peroxidase in Caulobacter crescentus. Infect. Immun. 181:6150–6150. Ravenscroft, N., S. G. Walker, G. G., Dutton, and J. Smith. 1992. Identification, isolation, and structural studies of the outer membrane lipopolysaccharide of Caulobacter crescentus. J. Bacteriol. 174:7595–7605. Ray, W. C. 1976. An assessment of investigations conducted in the USA on Brucella abortus strain 45/20 bacterins. Dev. Biol. Stand. 31:335–342. Ray, W. C. 1977. The epidemiology of Brucella abortus. In: R. M. Crawford and R. J. Hidalgo (Eds) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 102–115. Reisenauer, A., L. S. Kahng, S. McCollum, and L. Shapiro. 1999. Bacterial DNA methylation: A cell cycle regulator? J. Bacteriol. 181:5135–5139. Rennick, D. M., P. R. Morrow, and E. Benjamini. 1983. Functional heterogeneity of memory B lymphocytes. in vivo analysis of TD-primed B cells responsive to secondary stimulation with TD and TI antigens. J. Immunol. 131:561–566. Renoux, M., and G. Renoux. 1977. Brucellosis, immunodepression, and levamisole. Lancet 1:372. Reschke, D. K., M. E. Frazier, and L. P. Mallavia. 1990. Transformation of Rochalimaea quintana, a member of the family Rickettsiaceae. J. Bacteriol. 172:5130–5134. Rest, R. F., and D. C. Robertson. 1974. Glucose transport in Brucella abortus. J. Bacteriol. 118:250–258. Rest, R. F., and D. C. Robertson. 1975. Characterization of the electron transport system in Brucella abortus. J. Bacteriol. 122:139–144. Richardson, M., and J. N. Holt. 1964. Multiplication of Brucella in cultured lymphoid and non-lymphoid cells. J. Bacteriol. 88:1163–1168. Riezu-Boj, J. I., I. Moriyó n, J. M. Blasco, C. M. Marín, and R. Díaz. 1986. Comparison of lipopolysaccharide and outer membrane protein-lipopolysaccharide extracts in an enzyme-linked immunosorbent assay for the diagnosis of Brucella ovis infection. J. Clin. Microbiol. 23:938– 942. Riezu-Boj, J. I., I. Moriyó n, J. M. Blasco, C. Gamazo, and R. Díaz. 1990. Antibody response to Brucella ovis outer

450

E. Moreno and I. Moriyó n

membrane proteins in ovine brucellosis. Infect. Immun. 58:489–494. Rigby, C. E. 1990. The brucellaphages. In: K. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 121–129. Riglar, C., and C. Cheers. 1980. Macrophage activation during experimental murine brucellosis. II: Inhibition of in vitro lymphocyte proliferation by Brucella-activated macrophages. Cell. Immunol. 49:154–167. Riley, M., and A. Anilionis. 1978. Evolution of the bacterial genome. Ann. Rev. Microbiol. 32:519–560. Riley, L. K., and D. C. Robertson. 1984a. Brucellacidal activity of human and bovine polymorphonuclear leukocyte granule extracts against smooth and rough strains of Brucella abortus. Infect. Immun. 46:231–236. Riley L. K., and D. C. Robertson. 1984b. Ingestion an intracellular survival of Brucella abortus in human and bovine polymorphonuclear leukocytes. Infect. Immun. 46:224–230. Rivilla, R., and J. A. Downie. 1994. Identification of Rhizobium leguminosarum gene homologous to nodT but located outside the symbiotic plasmid. Gene 144:87–91. Robertson, D. C., and W. G. McCullough. 1968a. The glucose catabolism of the genus Brucella. I: Evaluation of pathways. Arch. Biochem. Biophys. 127:263–273. Robertson, D. C., and W. G. McCullough. 1968b. The glucose catabolism of the genus Brucella. II: Cell-free studies with B. abortus (S-19). Arch. Biochem. Biophys. 127:445–456. Robertson, L., I. D. Farrell, and P. M. Hinchliffe. 1977. The isolation of Brucellae from contaminated sources: A review. Br. Vet. J. 133:193–200. Robertson, G. T., and R. M. Roop 2nd. 1999. The Brucella abortus host factor I (HF-I) contributes to the stress resistance during stationary phase and is a major determinant of virulence in mice. Molec. Microbiol. 34:690– 700. Robertson, G. T., M. E. Kovach, C. A. Allen, T. A. Ficht, and R. M. Roop 2nd. 2000a. The Brucella abortus Lon functions as a generalized stress response protease and is required for wild-type virulence in BALB/c mice. Molec. Microbiol. 35:577–588. Robertson, G. T., A. Reisenauer, R. Wright, R. B. Jensen, A. Jensen, L. Shapiro, and R. M. Roop 2nd. 2000b. The Brucella abortus CcrM DNA methyltransferase is essential for viability, and its overexpression attenuates intracellular replication in macrophages. J. Bacteriol. 182:3482–3489. Rodríguez, M. C., A. Froger, J. P. Rolland, D. Thomas, J. Aguero, C. Delamarche, and J. M. García-Lobo. 2000. A functional water channel protein in the pathogenic bacterium Brucella abortus. Microbiology 146 :3251– 3257. Rodríguez-Torres, A., R. Landínez, and R. HernándezMejía. 1977. Diagnó stico bacterioló gico de la brucelosis humana. In: A. Rodríguez-Torres (Ed.) Guía Práctica de Diagnó stico de la Brucelosis Humana. Sever-Cuesta. Valladolid, Spain. 11–17. Roerink, J. H. G. 1969. Experience on the safety and effectiveness of 45/20 vaccine under field conditions. Vet. Rec. 85:269–270. Rojas, N., E. Freer, A. Weintraub, M. Ramirez, S. Lind, and E. Moreno. 1994. Immunochemical identification of Brucella abortus lipopolysaccharide epitopes. Clin. Diagn. Lab. Immunol. 1:206–213.

CHAPTER 3.1.16 Romero, C., C. Gamazo, M. Pardo, and I. Ló pez-Goñ i. 1995a. Specific detection of Brucella DNA by PCR. J. Clin. Microbiol. 33:615–617. , R. Díaz, J. M. Blasco, and Romero, C., M. Pardo, M. J. Grilló I. Ló pez-Goñ i. 1995b. Evaluation of PCR and indirect enzyme-linked immunosorbent assay on milk samples for diagnosis of brucellosis in dairy cattle. J. Clin. Microbiol. 33:3198–3200. Romero, C., and I. Ló pez-Goñ i. 1999. Improved method for purification of bacterial DNA from bovine milk for detection of Brucella spp. by PCR. Appl. Environ. Microbiol. 65:3735–3737. Roop, R. M., T. W. Fletcher, N. M. Sriranganathan, S. M. Boyle, and G. G. Schurig. 1994. Identification of an immunoreactive Brucella abortus HtrA stress response protein homolog. Infect. Immun. 62:1000–1007. Rossetti, O. L., A. I. Arese, M. L. Boschiroli, and S. L. Cravero. 1996. Cloning of Brucella abortus gene and characterization of expressed 26-kilodalton periplasmic protein: Potential use for diagnosis. J. Clin. Microbiol. 34:165–169. Rousseau, P., C. Ricard, and J. Cameron. 1987. Improvements in the fermenter growth of Brucella abortus strain 19. Appl. Microbiol. Biotechnol. 26:422–426. Roux, V., and D. Raoult. 1993. Genotypic identification and phylogenetic analysis of the spotted fever group rickettsiae by pulsed-field gel electrophoresis. J. Bacteriol. 175:4895–4904. Rubbi, C. P. 1991. Antígenos Proteicos de Brucella abortus: Aislamiento y Estudio de su Asociació n al LPS y de Factores que Afectan a la Producció n de Anticuerpos Monoclonales Contra los Mismos (PhD thesis). Facultad de Ciencias Exactas, Universidad Nacional de la Plata. La Plata , Argentina. 1–137. Rucavado, A., E. Moreno, and J. M. Gutiérrez. 1996. Effect of adjuvants on the antibody response of mice to Bothrops asper (Terciopelo) snake venom. Brazil. J. Med. Biol. Res. 29:1337–1340. Ruckerbauer, G. M., M. M. Garcia, C. E. Rigby, F. J. Robertson, B. S. Samagh, and B. W. Stemshorn. 1984. An hemolysis-in-gel test for bovine brucellosis. Dev. Biol. Stand. 56:513–520. Russa, R. Urbanik-Sypniewska; Lindstöm, and K; H. Mayer. 1995. Chemical Characterization of two lipopolysaccharuide species isolated from Rhizobium loti NZP2213. Arch. Microbiol. 163, :345–351. Russell, L. H. 1977. Epidemiology. In: R. M. Crawford and R. J. Hidalgo (Eds) Bovine Brucellosis. Texas A&M University Press. College Station, TX. 101–174. Sacks, T., and D. M. Kilnman. 1997. Long term efect of primary immunization on subsequent immune responsivness. Cell. Immunol. 177:162–168. Saegerman, C., T. K. Vo, W. L. De, D. Gilson, A. Bastin, G. Dubray, P. Flanagan, J. N. Limet, J. J. Letesson, and J. Godfroid. 1999. Diagnosis of bovine brucellosis by skin test: Conditions for the test and evaluation of its performance. Vet. Rec. 145:214–218. Saha, A. K., N. K. Mukhopadhyay, J. N. Dowling, T. A. Ficht, L. G. Adams, and R. H. Glew. 1990. Characterization of a phosphomonoesterase from Brucella abortus. Infect. Immun. 58:1153–1158. Saitou, N., and M. Nei. 1987. The neighbour-joining method: A new method for reconstructing phylogenetic trees. Molec. Biol. Evol. 4:406–425. Salih-Alj Debbarh, H., A. Cloeckaert, M. S. Zygmunt, and G. Dubray. 1995. Identification of sero-reactive Brucella

CHAPTER 3.1.16 melitensis cytosoluble proteins which discriminate between antibodies elicited by infection and Rev. 1 vaccination in sheep. Vet. Microbiol. 44:37–48. Salih-Alj Debbarh, H., M. S. Zygmunt, G. Dubray, and A. Cloeckaert. 1996. Competitive enzyme-linked immunosorbent assay using monoclonal antibodies to the Brucella melitensis BP26 protein to evaluate antibody responses in infected and B. melitensis Rev. 1 vaccinated sheep. Vet. Microbiol. 53:325–337. Salyers, A. A., and D. D. Whitt. 1994. Bacterial Pathogenesis: A Molecular Approach. American Society for Microbiology. Washington DC , 273–281. Samartino, L., D. Gall, R. Gregoret, and K. H. Nielsen. 1999. Validation of enzyme-linked immunosorbent assays for the diagnosis of bovine brucellosis. Vet. Microbiol. 70:193–200. Samartino, L. E., M. Fort, R. Gregoret, G. G. Schurig. 2000. Use of Brucella abortus vaccine strain RB51 in pregnant cows after calfhood vaccination with 19 in Argentina. Prev. Vet. Med. 45:193–199. n, Sánchez, D. O., R. O. Zandomeni, S. Cravero, R. E. Verdú E. Pierrou, P. Faccio, G. Díaz, S. Lazavecchia, A. C. C. Frasch, S. G. E. Andersson, O. L. Rossetti, O. Grau, and R. A. Ugalde. 2001. Gene discovery through sequencing of Brucella abortus. Infect. Immun. 69:865–868. Sangari, F. J. 1993. Caracterizació n de la Regió n eri de Brucella abortus: Su Aplicació n al Diagnó stico Molecular (PhD thesis). University of Cantabria. Santander, Spain. 1–98. Sangari, F. J., and J. Agü ero. 1994a. Identification of Brucella abortus B19 vaccine strain by the detection of DNA polymorphism at the ery locus. Vaccine 12:435–438. Sangari, F. J., J. M. García-Lobo, J. Agü ero. 1994b. The Brucella abortus vaccine strain S19 carries a deletion in the erythritol catabolic genes. FEMS Microbiol. Lett. 121:337–342. Sangari, F. J., J. Díaz, A. Seoane, J. Agü ero, M. J. Grilló , and J. M. Garcí-Lobo. 2000. The role of urease in Brucella infection. In: D. O’Callaghan (Ed.) Brucellosis 2000. INSERM, Facultéde Médicine. Nîmes, France. 97:79– 80. Sangari, F. J., J. Aguero, and J. M. García-Lobo. 2000. The genes for erythritol catabolism are organized as an inducible operon in Brucella abortus. Microbiology 146:487–495. Sano, Y., S. Kondo, Y. Isshiki, T. Shimada, and K. Hisatsune. 1996. An N-[(R)-(-)-2-hydroxypropionyl]-alpha-Lperosamine homopolymer constitutes the O polysaccharide chain of the lipopolysaccharide from Vibrio cholerae O144 which has antigenic factor(s) in common with V. cholerae O76. Microbiol. Immunol. 40:735–741. Sarmento, A., and R. Appelberg. 1996. Involvement of reactive oxygen intermediates in tumor necrosis factor alpha-dependent bacteriostasis of Mycobacterium avium. Infect. Immun. 64:3224–3230. Scherer, D. C., I. DeBuron-Connors, and M. F. Minnick. 1993. Characterization of Bartonella bacilliformis flagella and effect of antiflagellin antibodies on invasion of human erythrocytes. Infect. Immun. 61:4962–4971. Schmiederer, M., R. Arcenas, R. Widen, N. Valkov, and B. Anderson. 2001. Intracellular induction of the Bartonella henselae virB operon by human endothelial cells. Infect. Immun. 69:6495–6502. Schmitt-Slomska, J., R. Caravano, M. Anoal, B. Gay, and J. Roux. 1981. Isolation of L-forms from the spleens of

The Genus Brucella

451

Brucella suis-infected, penicillin-treated mice. Ann. Microbiol. Paris 132:253–265. Schoenlein P. V., and B. Ely. 1983. Plamids and bacteriocins in Caulobacter species. J. Bacteriol. 153:1092–1094. Schoerner, C., K. Wartenberg, and M. Rollinghoff. 1990. Differentiation of serological responses to Yersinia enterocolitica serotype O9 and Brucella species by immunoblot or enzyme-linked immunosorbent assay using whole bacteria and Yersinia outer membrane proteins. J. Clin. Microbiol. 28:1570–1574. Schorey, J. S., M. C. Carrol, E. J. Brown. 1997. A macrophage invasion mechanism of pathogenic mycobacteria. Science 277:1091–1093. Schultz, R. M., M. A. Chirigos, N. A. Pavlidis, J. S. Youngner. 1978. Macrophage activation and antitumor activity of a Brucella abortus ether extract, Bru-Pel. Cancer Treat. Rep. 62:1937–1941. Schurig, G. G., A. T. Pringle, and S. S. Breese. 1981. Localization of brucella antigens that elicit a humoral immune response in Brucella abortus-infected cells. Infect. Immun. 34:1000–1007. Schurig, G. G., R. M. Roop, T. Bagchi, S. Boyle, D. Buhrman, and N. M. Sriranganathan. 1991. Biological properties of RB51; a stable rough strain of Brucella abortus. Vet. Microbiol. 28:171–188. Schwedock, J., and S. R. Long. 1994. An open reading frame downstream of Rhizobium meliloti nod Q1 shows nucleotide sequence similarity to an Agrobacterium tumefaciens insertion sequence. Molec. Plant-Microbe Interact. 7:151–153. Scianimanico, S., M. Desrosiers, J. F. Dermine, S. Méresse, A. Descoteaux, and M. Desjardins. 1999. Impaired recruitment of the small GTPase rab7 correlates with the inhibition of phagosome maturation by Leishmania donovani promastigotes. Cell. Microbiol. 1:19–32. Scott, L. M., C. I. Civin, P. Rorth, and A. D. Friedman. 1992. A novel temporal expression pattern of three C/EBP family members in differentiating myelomonocytic cells. Blood 80:1725–1735. Serikawa, T. H. Takada, Y. Kondo, T. Muraguchi, and J. Yamada. 1984. Multiplication of Brucella suis in male reproductive organs and detection of autoantibody to spermatozoa in canine brucellosis. In: L. Vallete and W. Heunessen (Eds.) Biological Standardization: 3rd International Symposium on Brucellosis. Kerger. Basle, Switzerland. 56:295. Serre, A., J. P. Vendrell, M. F. Huguet, and A. Cannat. 1982. In vitro studies on the adjuvanticity of Brucella fractions. Infect. Immun. 38:413–418. Sha, Z., T. J. Stabel, and J. E. Mayfield. 1994. Brucella abortus catalase is a periplasmic protein lacking a standard signal sequence. J. Bacteriol. 176:7375–7377. Shuterland, S. S., and J. Searson. 1990. The immune response to Brucella abortus. In: K. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 65–82. Sieira, R., and D. J. Comerci, D. O. Sánchez, R. A. Ugalde. 2000. A homologue of an operon required for DNA transfer in Agrobacterium is required in Brucella abortus for virulence and intracellular multiplication. Infect. Immun. 182:4849–4655. Sifuentes-Rincó n, A. M., A. Revol, and H. A. BarreraSaldana. 1997. Detection and differentiation of the six Brucella species by polymerase chain reaction. Molec. Med. 3:734–739.

452

E. Moreno and I. Moriyó n

Silverstein, S. C., and T. H. Steinberg. 1990. Host defense against bacterial and fungal infections. In: D. B. Davies, R. Dulbecco, H. N. Eisen, and H. S. Ginsberg (Eds.) Microbiology, 4th ed. J. B. Lippincott. Philadelphia, PA. 485–505. Simon, S. M., and G. Blobel. 1991. A protein-conducting channel in the endoplasmic reticulum. Cell 65:371–380. Sinai, A. P., and K. A. Joiner. 1997a. The cell biology of nonfusogenic pathogen vacuoles. Ann. Rev. Microbiol. 51:415–462. Sinai, A. P., P. Webster, and K. A. Joiner. 1997b. Association of host cell endoplasmic reticulum and mitochondria with the Toxoplasma gondii parasitophorous vacuole membrane: A high affinity interaction. J. Cell Sci. 110:2117–2128. Sly, L. I., T. L. Cox, and T. B. Beckenham. 1999. The phylogenetic relationships of Caulobacter, Asticcacaulis and Brevundimonas species and their taxonomic implications. Int. J. Syst. Bacteriol. 49:483–288. Smith, H., J. D. Anderson, and J. Keppie. 1965. The inhibition of growth of brucellas in vitro and in vivo by analogues of erythritol. J. Gen. Microbiol. 38:101–108. Sneath, P. H. A. 1984a. Bacterial nomenclature. In: N. R. Krieg and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. William and Wilkins. Baltimore, MD. 1:19–23. Sneath, P. H. A. 1984b. Numerical taxonomy. In: N. R. Krieg and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. William and Wilkins. Baltimore, MD. 1:5– 7. Sniegowski, P. 1997. Evolution: Setting the mutation rate. Curr. Biol. 7:R487–R488. So, R. B., J. K. Ladha, and J. P. W. Young. 1994. Photosynthetic symbionts of Aeschynomene spp. form a cluster with Bradyrhizobia on the basis of fatty acid and rRNA analysis. Int. J. Syst. Bacteriol. 44:392–403. Sola-Landa, A., J. Pizarro-Cerdá, J. M. Grilló , E. Moreno, I. Moriyó n, J. M. Blasco, J.-P. Gorvel, and I. Ló pez-Goñ i. 1998. A two component regulatory system playing a critical role in plant pathogens and endosymbionts is present in Brucella abortus and controls cell invasion and virulence. Molec. Microbiol. 29:5265–5273. Soto, L., X. Rojas, and O. Alonso. 1991. Estudio in vitro del efecto de fracciones de pared celular de Brucella sobre la actividad de leucocitos polimorfonucleares bovinos. Arch. Med. Vet. 23:27–33. Southern Jr., P. M. 1996. Bacteremia due to Agrobacterium tumefaciens (radiobacter) report of infection in a pregnant women and her stillborn fetus. Diagn. Microbiol. Infect. Dis. 24:43–45. Spencer, T. L., and G. W. Burgess. 1984. Enzyme-linked immunosorbent assay for Brucella ovis specific antibody in ram sera. Res. Vet. Sci. 36:194–198. Spencer, S. A., E. S. Broughton, S. Hamid, and D. B. Young. 1994. Immunoblot studies in the differential diagnosis of porcine brucellosis: An immunodominant 62 kDa protein is related to the mycobacterial 65 kDa heat shock protein (HSP-65). Vet. Microbiol. 39:47–60. Sperry, J. F., and D. C. Robertson. 1975. Erythritol catabolism by Brucella abortus. J. Bacteriol. 121:619–630. Spink, W. W. 1956. The Nature of Brucellosis. University of Minnesota Press. Minneapolis, MN. 1–420. Splitter, G. A., and K. M. Everlith. 1986. Collaboration of bovine T lymphocytes and macrophages in T-lymphocyte response to Brusella abortus. Infect. Immun. 51:776–783.

CHAPTER 3.1.16 Splitter, G. A., and K. M. Everlith. 1989. Brusella abortus regulates bovine macrophage-T-cell interaction by major histocompatibility complex class II and interleukin-1 expression. Infect. Immun. 57:1151–1157. Stabel, T. J., Z. Sha, and J. E. Mayfield. 1994. Periplasmic location of Brucella abortus Cu/Zn superoxide dismutase. Vet. Microbiol. 38:307–314. Stackebrandt, E., and B. M. Goebel. 1994. A place for DNADNA reassociation and 16S rRNA sequence analysis in the present species definition in bacteriology. Int. J. Syst. Bacteriol. 44:846–849. Stackebrandt, E., A. Fischer, T. Roggentin, U. Wehmeyer, D. Bomar, and J. Smida. 1988. A phylogenetic survey of budding, and/or prosthecate, non-phototrophic eubacteria: membership of Hyphomicrobium, Hyphomonas, Pedomicrobium, Filomicrobium,Caulobacter and “Dichotomicrobium” to the alpha-subdivision of purple non-sulfur bacteria. Arch. Microbiol. 149:547– 556. Staley, J. T., and N. R. Krieg. 1984. Classification of procaryotic organisms: An overview. In: N. R. Krieg and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. William and Wilkins. Baltimore, MD. 1:1– 4. Stang, E., J. Kartenbeck, and R. G. Parton. 1997. Major histocompatibility complex class I molecules mediate association of SV40 with caveolae. Molec. Biol. Cell. 8:47– 57. Steele-Mortimer, O., S. Méresse, J. P. Gorvel, B. H. Toh, and B. B. Finlay. 1999. Biogenesis of Salmonella typhimurium-containing vacuoles in epithelial cells involves interactions with the early endocytic pathway. Cell. Microbiol. 1:33–49. Steinman, H. M., and B. Ely. 1990. Copper-zinc superoxide dismutase of Caulobacter crescentus: cloning, sequencing, and mapping of the gene and periplasmic location of the enzyme. J. Bacteriol. 172:2901–2910. Sterba, F. 1984. Morphological changes in brucellosisin guines pigs due to experimental infection with Brucella suis. Vet. Med. (Praha) 29:353–358. Stevens, M. G., S. G. Hennager, S. C. Olsen, and N. F. Cheville. 1994a. Serologic responses in diagnostic tests for brucellosis in cattle vaccinated with Brucella abortus 19 or RB51. J. Clin. Microbiol. 32:1065–1066. Stevens, M. G., S. C. Olsen, G. W. Pugh Jr., and M. V. Palmer. 1994b. Immune and pathologic responses in mice infected with Brucella abortus 19, RB51, or 2308. Infect. Immun. 62:3206–3212. Suárez, C. E., G. A. Pacheco, and A. M. Vigliocco. 1990. Immunochemical studies of oligosaccharides obtained from the lipopolysaccharide of Brucella ovis. Vet. Microbiol. 22:329–334. Sun L. V., J. M. Foster, G. Tzertzinis, M. Ono, C. Bandi, B. E. Slatko, and S. L. O’Neill. 2001. Determination of Wolbachia genome size by pulsed-field gel electrophoresis. J. Bacteriol. 183:2219–25. Sutra, L., J. P. Caffin, and G. Dubray. 1986. Role of milk immunoglobulins in the Brucella milk ring test. Vet. Microbiol. 12:359–366. Suwanto, A., and S. Kaplan. 1992. Chromosome transfer in Rhodobacter sphaeroides: Hfr formation and genetic evidence for two unique circular chromosomes. J. Bacteriol. 174:1135–1145. Swanson, M. S., and R. R. Isberg. 1995. Association of Legionella pneumophila with the macrophage endoplasmic reticulum. Infect. Immun. 63:3609–3620.

CHAPTER 3.1.16 Syvanen, M. 1994. Horizontal gene transfer: Evidence and possible consequences. Ann. Rev. Genet. 28:237–261. Tabatabai, L. B., and S. G. Hennager. 1994. Cattle serologically positive for Brucella abortus have antibodies to B. abortus Cu-Zn superoxide dismutase. Clin. Diagn. Lab. Immunol. 1:506–510. Taminiau B., M. Daykin, S. Swift, M. L. Boschiroli, A. Tibor, P. Lestrate, X. De Bolle, D. O’Callaghan, P. Williams, and J. J. Letesson. 2002. Identification of a quorumsensing signal molecule in the facultative intracellular pathogen Brucella melitensis. Infect. Immun. 70:3004– 11. Tanaka, S., T. Suto, Y. Isayama, R. Azuma, and H. Hatakeyama. 1977. Chemo-taxonomical studies on fatty acids of “Brucella” species. Ann. Sclavo 19:67–82. Tanaka., T., S. Akira, K. Yoshida, M. Umemoto, Y. Yoneda, N. Shirafuji, H. Fujiwara, S. Suematsu, N. Yoshida, and T. Kishimoto. 1995. Targeted disruption of the NF-IL6 gene discloses its essential role in bacteria killing and tumor cytotoxicity by macrophages. Cell 80:353–361. Taran, I. F., and N. A. Rybasov. 1972. Comparative study of susceptibility and sensitivity to infection of laboratory animals and sheep with various species of Brucella. Zh. Mikrobilo. Epidemid. Immunobiol. 10:97–108. Tardieux, I., P. Webster, J. Ravesloot, W. Boron, J. A. Lunn, H. E. Heuser, and N. W. Andrews. 1992. Lysosome recruitment and fusion are early events required for trypanosome invasion of mammalian cells. Cell 71:1117– 1130. Tatum, F. M., P. G. Detilleux, J. M. Sacks, and S. M. Halling. 1992. Construction of Cu-Zn superoxide dismutase deletion mutants of Brucella abortus: Analysis of survival in vitro in epithelial and phagocytic cells and in vivo in mice. Infect. Immun. 60:2863–2869. Tatum, F. M., D. C. Morfitt, and S. M. Halling. 1993. Construction of a Brucella abortus RecA mutant and its survival in mice. Microb. Pathog. 14:177–185. Tatum, F. M., N. F. Cheville, and D. C. Morfitt. 1994. Cloning, characterization and construction of htrA and htrA-like mutants of Brucella abortus and their survival in BALB/ c mice. Microb. Pathog. 17:23–36. Tcherneva, E., N. Rijpens, B. Jersek, and L. M. Herman. 2000. Differentiation of Brucella species by random amplified polymorphic DNA analysis. J. Appl. Microbiol. 88:68–80. Teixeira-Gomes, A. P., A. Cloeckaert, G. Bezard, R. A. Bowden, G. Dubray, and M. S. Zygmunt. 1997. Identification and characterization of Brucella ovis immunogenic proteins using two-dimensional electrophoresis and immunoblotting. Electrophoresis 18:1491–1497. Teixeira-Gomes, A. P., A. Cloeckaert, and M. S. Zygnunt. 2000. Characterization of heat, oxidative and acid stress responses in Brucella melitensis. Infect. Immun. 68:2954–2961. Templeton, J. W., and L. G. Adams. 1990. Natural resistance to bovine brucellosis. In: L. G. Adams (Ed.) Advances in Brucellosis Research. Texas A&M University Press. College Station, TX. 144–150. Tempst, P., and J. Van Beeumen. 1983. The complete aminoacid sequence of the low-spin class II cytochrome c-556 from Agrobacterium tumefaciens strain B2a. Eur. J. Biochem. 129:603–614. Tenay, A., and S. Strober. 1985. T cell regulation of the thymus-independent antibody response to trinitrophenylated-Brucella abortus (TNP-BA). J. Immunol. 134:3669–3674.

The Genus Brucella

453

Teodorescu, M., E. P. Mayer, and S. Day. 1977. Enumeration and identification of humanleukemic lymphocytes by their natural binding of bacteria. Cancer. Res. 37:1715– 1718. Thiele, O. W., and W. Kher. 1969. Die “freien” Lipide aus Brucella abortus Bang. Eur. J. Biochem. 9:167–165. Thiele, O. W., and G. Schwinn. 1973. The free lipids of Brucella melitensis and Bordetella pertussis. Eur. J. Biochem. 34:333–334. Thoen, C. O., D. E. Pietz, A. L. Armbrust, and R. Harrington. 1979. Enzyme immunoassay for detecting Brucella antibodies in cow’s milk. J. Clin. Microbiol. 10:222–225. Tibor, A., B. Decelle, and J. J. Letesson. 1999. Outer membrane proteins Omp10, Omp16, and Omp19 of Brucella spp. are lipoproteins. Infect. Immun. 67:4960– 4962. Tighe, S. W., P. deLajudie, K. Dipietro, L. Lindstrom, G. Nick, and B. D. Jarvis. 2000. Bradyrhizobium, Mesorhizobium, Rhizobium ancl Sinorhizobium species using the Shelock Microbial Identification System. Int. J. Syst. E. Microbiol. 2:787–801. Tobias L., D. O. Cordes, and G. G. Schurig. 1993. Placental pathology of the pregnant mouse inoculated with Brucella abortus strain 2308. Vet. Pathol. 30:119–129. Trap, D., and R. Gaumont. 1982. Comparaison entre electrosynérèse et épreuves sérologiques classiques dans le diagnostic de la brucellose ovine. Ann. Rech. Vet. 13:33– 39. Uchiyama, T., Y. Kamagata, and M. Yoshioka. 1984. Mechanism of lipopolysaccharide-induced immunosuppression: Immunological activity of B cell subsets responding to T-dependent or T independent antigens in lipopolysaccharide-preinjected mice. Infect. Immun. 45:367–371. Ugalde, R. A. 1999. Intracellular lifestyle of Brucella spp. Common genes with other animal pathogens, plant pathogens, and endosymbionts. Microb. Infect. 1:1211– 1219. Ugalde, J. E., C. Czibener, M. F. Feldman, and R. A. Ugalde. 2000. Indentification and characterization of the Brucella abortus phosphoglucomutase gene: Role of lipopolysacchride in virulence and intracellular multiplication. Infect. Immun. 10:5716–5723. Urbanik-Sypniewska, T., A. Choma, J. Kutkowska, T. Kaminska, M. Kandefer-Szerszen, R. Russa, and J. Dolecka. 2000. Cytokine induction activities of rhizobial and mesorhizobial lipopolysaccharides of different toxicity. Immunobiology 202:408–420. Uza, F. A., L. Samartino, G. G. Schurig, A. Carrasco, K. H. Nielsen, R. F. Cabrera, and H. R. Taddeo. 2000. Effect of vaccination with Brucella abortus strain RB51 on heifers and pregnant cattle. Vet. Res. Commun. 24:143– 151. Vaara, M. 1992. Agents that increase the permeability of the outer membrane. Microbiol. Rev. 56:395–411. Valitutti, S., S. Muller, M. Cella, E. Padovan, and A. Lanzavecchia. 1995. Serial triggering of many T-cell receptors by a few peptide-MHC complexes. Nature 375:148–151. Van Nhieu, G. T., and P. J. Sansonetti. 2000. Cell adhesion molecules and bacterial pathogens. In: P. Cossart, P. Bouquet, S. Normark, and R. Rappuoli (Eds.) Cellular Microbiology. American Society for Microbiology. Washington DC, 97–111. Vanzini, V. R., N. Aguirre, C. I. Lugaresi, S. T. de Echaide, V. G. de Canavesio, A. A. Guglielmone, M. D.

454

E. Moreno and I. Moriyó n

Marchesino, and K. H. Nielsen. 1998. Evaluation of an indirect ELISA for the diagnosis of bovine brucellosis in milk and serum samples in dairy cattle in Argentina. Prev. Vet. Med. 36:211–217. Velasco, J., H. Moll, E. V. Vinogradov, I. Moriyó n, and U. Zähringer. 1996. Determination of the O-specific polysaccharide structure in the lipopolysaccharide of Ochrobactrum anthropi LMG 3331. Carbohydr. Res. 287:123–126. Velasco, J., R. Díaz, M. J. Grilló , M. Barberán, C. M. Marín, J. M. Blasco, and I. Moriyó n. 1997. Antibody and delayed-type hypersensitivity responses to Ochrobactrum anthropi cytosolic and outer membrane antigens in infections by smooth and rough Brucella spp. Clin. Diagn. Lab. Immunol. 4:279–284. Velasco, J., H. Moll, Y. A. Knirel, V. Sinnwell, I. Moriyó n, and U. Zähringer. 1998a. Structural studies on the lipopolysaccharide from a rough strain of Ochrobactrum anthropi containing a 2,3-diamino-2,3-dideoxy-Dglucose disaccharide lipid A backbone. Carbohydr. Res. 306:283–290. pez-Goñ i, J. Leiva, R. Díaz, and Velasco, J., C. Romero, I. Ló I. Moriyó n. 1998b. Evaluation of the relatedness of Brucella spp. and Ochrobactrum antrophi and description of Ochrobactrum intermedium sp. nov., a new species with close relationship to Brucella spp. Int. J. Syst. Bacteriol. 48:759–768. Velasco, J., J. A. Bengoechea, K. Brandenburg, B. Lindner, U. Seydel, D. Gonzalez, U. Zahringer, E. Moreno, and I. Moriyó n. 2000. Brucella abortus and its closest phylogenetic relative, Ochrobactrum spp., differ in outer membrane permeability and cationic peptide resistance. Infect. Immun. 68:3210–3218. Vemulapalli, R., J. R. McQuiston, G. G. Schurig, N. Sriranganathan, S. M. Halling, and S. M. Boyle. 1999. Identification of and IS711 element interrupting the wboA gene of Brucella abortus vaccine strain RB51 and a PCR assay to distinguish strain RB51 from other Brucella species and strains. Clin. Diagn. Lab. Immunol. 6:760– 764. Vemulapalli, R., S. M. Yhe-Boyle., N. Sriranganathan, and G. G. Schurig. 2000. Institution Brucella abortus strain RB51 as a vector for heterologous protein expression and induction of specific Th1 type immune responses. Infect. Immun. 68:3290–3296. Verger, J., F. Grimont, P. A. D. Grimont, and M. Grayon. 1985. Brucella, a monospecific genus as shown by deoxyribonucleic acid hybridization. Int. J. Syst. Bacteriol. 35:292–295. Verger, J. M., M. Grayon, E. Zundel, P. Lechopier, and V. Olivier-Bernardin. 1995. Comparison of the efficacy of Brucella suis strain 2 and Brucella melitensis Rev. 1 live vaccines against a Brucella melitensis experimental infection in pregnant ewes. Vaccine 13:191–196. Verger, J. M., M. Grayon, A. Tibor, V. Wansard, J. J. Letesson, and A. Cloeckaert. 1998. Differentiation of Brucella melitensis, B. ovis and B. suis biovar 2 strains by use of membrane protein- or cytoplasmic protein-specific gene probes. Res. Microbiol. 149:509–517. Via, L. E., D. Deretic, R. J. Ulmer, N. S. Hible, L. A. Huber, and V. Deretic. 1997. Arrest of mycobacterial phagosome maturation is caused by a block in vesicle fusion between stages. controlled by rab5 and rab7. J. Biol. Chem. 272:13326–13331. Viale, A. M., A. K. Arakaki, F. C. Soncini, and R. G. Ferreyra. 1994. Evolutionary relationships among eubacterial

CHAPTER 3.1.16 groups as inferred from GroEL (chaperonin) sequence comparisons. Int. J. Syst. Bacteriol. 44:527–533. Vigliocco, A. M., P. S. Silva-Paulo, J. Mestre, G. C. Briones, G. Draghi, M. Tossi, and K. H. Nielsen. 1997. Development and validation of an indirect enzyme immunoassay for detection of ovine antibody to Brucella ovis. Vet. Microbiol. 54:357–368. Viola, A., and A. Lanzavecchia. 1996. T cell activation determined by T cell receptor number and tunable thresholds. Science 273:104–106. Vitas, A. I., R. Díaz, and C. Gamazo. 1997. Protective effect of liposomal gentamicin against systemic acute murine brucellosis. Chemotherapy 43:204–210. Vizcaíno, N., A. Cloeckaert, G. Dubray, and M. S. Zygmunt. 1996. Cloning, nucleotide sequence, and expression of the gene coding for a ribosome releasing factorhomologous protein of Brucella melitensis. Infect. Immun. 64:4834–4837. Vizcaíno, N., J. M. Verger, M. Grayon, M. S. Zygmunt, and A. Cloeckaert. 1997. DNA polymorphism at the omp-31 locus of Brucella spp.: Evidence for a large deletion in Brucella abortus, and other species-specific markers. Microbiology 143:2913–2921. Vizcaíno, N., A. Cloeckaert, M. S. Zygmunt, and L. Fernandez-Lago. 1999. Molecular characterization of Brucella species large DNA fragment deleted in Brucella abortus strains: Evidence for a locus involved in the synthesis of polysaccharide. Infect. Immun. 67:2700–2712. Vizcaíno, N., A. Cloeckaert, J. Verger, M. Grayon, and L. L. Fernandez. 2000. DNA polymorphism in the genus Brucella. Microb. Infect. 2:1089–1100. Vodovotz, Y., D. Russell, Q. W. Xie, C. Bogdan, and C. Nathan. 1995. Vesicle membrane association of nitric oxide synthase in primary mouse macrophages. J. Immunol. 154:2914–2925. Vysotskii, V. V., D. S. Kurdina, and N. N. Ostrovskaia. 1967. Ul’trastruktura brutsell. II: Ul’trastruktura R-kletok. Zh. Mikrobiol. Epidemiol. Immunobiol. 44:19–22. Vysotskii, V. V. 1968. Ul’trastruktyra brutsell. V. Protsess idssotsiatsii. Zh. Mikrobiol. Epidemiol. Immunobiol. 45:42–47. Watarai, M., S. Makino, Y. Fuji, K. Okamoto, and T. Shirahata. 2002. Modulation of Brucella-induced macropinocytosis by lipid rafts mediates intracellular replication. Cell. Microbiol. 46:341–356. Weber, A., H. G. Schiefer, and H. Krauss. 1977. Elektronenmikroskopische Darstellung anionisch geladener Gruppen auf der Zelloberflache von Brucella canis. Zentralbl. Bakteriol., Orig. A. 239:365–374. Weber, A., H. G. Schiefer, and H. Krauss. 1978. Elektronenmikroskopische Differenzierung der S- und R-Formen von Brucellen durch Behandlung mit polykationischem Ferritin. Zentralbl. Veterinarmed. B 25:324–328. Weisburg, W. G., M. E. Dobson, J. E. Samuel, G. A. Dasch, L. P. Mallavia, O. Baca, L. Mandelo, J. E. Sechrest, E. Weiss, and C. R. Woese. 1989. Phylogenetic diversity of rickettsiae. J. Bacteriol. 171:4202–4206. Weisburg, W. G., S. M. Barns, D. A. Pelletier, and D. J. Lane. 1991. 16S ribosomal DNA amplification of phylogenetic study. J. Bacteriol. 173:697–703. Weiss, E., and J. W. Moulder. 1984. Tribe 1: Rickettsiaceae Philip, 1953, 486AL. In: N. R. Krieg and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. William and Wilkins. Baltimore, MD. 1:688–701. Werren, J. H., and J. D. Bartos. 2001. Recombination in Wolbachia. Curr. Biol. 20:431–435.

CHAPTER 3.1.16 Weynants, V. E., J. Godfroid, B. Limbourg, C. Saegerman, and J. J. Letesson. 1995. Specific bovine brucellosis diagnosis based on in vitro antigen-specific gamma interferon production. J. Clin. Microbiol. 33:706–712. Weynants, V. E., D. Gilson, A. Cloeckaert, P. A. Denoel, A. Tibor, P. Thiange, J. N. Limet, and J. J. Letesson. 1996a. Characterization of a monoclonal antibody specific for Brucella smooth lipopolysaccharide and development of a competitive enzyme-linked immunosorbent assay to improve the serological diagnosis of brucellosis. Clin. Diagn. Lab. Immunol. 3:309–314. Weynants, V. E., A. Tibor, P. A. Denoel, C. Saegerman, J. Godfroid, P. Thiange, and J. J. Letesson. 1996b. Infection of cattle with Yersinia enterocolitica O:9 a cause of the false positive serological reactions in bovine brucellosis diagnostic tests. Vet. Microbiol. 48:101– 112. Weynants, V. E., D. Gilson, A. Cloeckaert, A. Tibor, P. A. Denoel, F. Godfroid, J. N. Limet, and J. J. Letesson. 1997. Characterization of smooth lipopolysaccharides and O polysaccharides of Brucella species by competition binding assays with monoclonal antibodies. Infect. Immun. 65:1939–1943. Whitfield, C., P. A. Amor, and R. Köplin. 1997. Modulation of the surface architecture of gram negative bacteria by the action of surface polymer:lipid A-core ligase and by determinants of polymer chain length. Molec. Microbiol. 23:629–638. Whitney, J. A., M. Gomez, D. Sheff, T. E. Kreis, and I. Mellman. 1995. Cytoplasmic coat proteins involved in endosome function. Cell 83:703–713. Williams, E. E., H. Rand, S. Rand, and R. J. O’Hara. 1995. A computer approach to the comparison of species in difficult taxonomic groups. Breviora 502:1–47. Wilson, J. B., and B. L. Dasinger. 1960. Biochemical properties of virulent and avirulent strains of brucellae. Proc. NY Acad. Sci. 88:1155–1166. Woese, C. R. 1994. There must be a prokaryote somewhere: Microbiology’s search for itself. Microbiol. Rev. 58:1–9. Wong, J. D., J. M. Janda, and P. S. Duffey. 1992. Preliminary studies on the use of carbon substrate utilization patterns for the identification of Brucella species. Diagn. Microbiol. Infect. Dis. 15:109–113. Wong, F. Y. K., E. Stackebrandt, J. K. Ladha, D. E. Fleischman, R. A. Date, and J. A. Fuerst. 1994. Phylogenetic analysis of Bradyrhizobium japonicum and photosynthetic stem-nodulating bacteria from Aeschynomene species grown in separated geographical regions. Appl. Environ. Microbiol. 60:940–946. Wrathall, A. E., E. S. Broughton, K. P. Gill, and G. P. Goldsmith. 1993. Serological reactions to Brucella species in British pigs. Vet. Rec. 132:449–454. Wright, S. D., and S. C. Silverstein. 1983. Receptors for C3b and C3bi promote phagocytosis but not the release of toxic oxygen from human phagocytes. J. Exp. Med. 158:2016–2023. Wright, P. F., and K. H. Nielsen. 1990a. Current and future serological methods. In: L. G. Adams (Ed.) Advances in Brucellosis Research. Texas A&M University Press. College Station, TX. 303–320. Wright, P. F., K. H. Nielsen, and W. A. Kelly. 1990b. Primary binding techniques for the serolodiagnosis of bovine brucellosis. In: K. H. Nielsen and J. R. Duncan (Eds.) Animal Brucellosis. CRC Press. Boca Raton, FL. 199– 226.

The Genus Brucella

455

Wright, R., C. Stephens, and L. Shapiro. 1997. The CcrM DNA methyltransferase is widespread in the alpha subdivision of Proteobacteria, and its essential functions are conserved in Rhizobium meliloti and Caulobacter crescentus. J. Bacteriol. 179:5869–5877. Xin, X. 1986. Orally administrable brucellosis vaccine: Brucella suis strain 2 vaccine. Vaccine 4:212–216. Yagupsky, P., N. Peled, J. Press, M. Abu Rashid, and O. Abramson. 1997. Rapid detection of Brucella melitensis from blood cultures by a commercial system. Eur. J. Clin. Microbiol. Infect. Dis. 16:605–607. Yanagi, M., and K. Yamasato. 1993. Phylogenetic analysis of the family Rhizobiaceae and related bacteria by sequencing of 16S rRNA gene using PCR and DNA sequencer. FEMS Microbiol. Lett. 107:115–120. Yantorno, O. Y., L. A. Mazza, A. P. Balatti, and W. Aguirre. 1978. Influencia de la técnica de esterilizació n y de los componentes del medio de cultivo en el desarrollo y disociació n de Brucella abortus cepa 19 en proceso sumergido. Rev. Asoc. Argent. Microbiol. 10:83–93. Young, E. J., C. I. Gomez, D. H. Yawn, D. M. Musher. 1979. Comparison of Brucella abortus and Brucella melitensis infections in mice and their effect on acquired cellular resistance. Infect. Immun. 26:680–685. Young, E. J. 1983. Human brucellosis. Rev. Infect. Dis. 5:821– 842. Young, E. J., M. Borchert, F. L. Kretzer, and D. M. Musher. 1985. Phagocytosis and killing of Brucella by human polymorphonuclear leukocytes. J. Infect. Dis. 151:682– 690. Young, J. P. W., and Wexler, M. 1988. Sym plasmid and chromosomal genotypes are correlated in field populations of Rhizobium leguminosarum. J. Gen. Microbiol. 134:2731–2739. Yuan, L., S. Inoue, Y. Saito, and O. Nakajima. 1993. An evaluation of the effects of cytokines on intracellular oxidative production in normal neutrophils by flow cytometry. Exp. Cell Res. 209:375–381. Zähringer, U., Y. A. Knirel, B. Lindner, J. H. Helbig, A. Sonesson, R. Marre, and E. T. Rietschel. 1995. The lipopolysaccharide of Legionella pneumophila serogroup 1 (strain Philadelphia 1): Chemical structure and biological significance. Prog. Clin. Res. 392:113– 139. Zambryski, P. 1988. Basic processes underlying agrobacteriamediated DNA transfer to plant cells. Ann. Rev. Genet. 22:1–30. Zezerov, E. G., V. S. Loginov, and A. S. Berezneva. 1985. Polypetide and phospholipid compositon of Rickettsia prowaseki membrane and its immunogenic properties. Zhurnal. Mikrobiol. Epidemiol. Immunobiol. Russia. 6:6–13. Zhan, Y., and C. Cheers. 1995. Differential induction of macrophage-derived cytokines by live and dead intracellular bacteria in vitro. Infect. Immun. 63:720–723. Zhan, Y., Z. Liu, and C. Cheers. 1996. Tumor necrosis factor alpha and interleukin-12 contribute to resistance to intracellular bacterium Brucella abortus by different mechanisms. Infect. Immun. 64:2782–2786. Zhang, J. 1992. A study on the role of immunosuppression in the pathogenesis of brucellosis. Acta Acad. Med. Sinicae 14:168–172. Zhang, J., B. Gao, C. Cun, X. Lu, H. Wang, X. Chen, and L. Tang. 1993. Immunosuppression in murine brucellosis. Chin. Med. Sci. J. 8:134–138.

456

E. Moreno and I. Moriyó n

Zimmerman, S. J., S. Gillikin, N. Sofat, W. R. Bartholomew, and D. Amsterdam. 1990. Case report and seeded blood culture study of Brucella bacteremia. J. Clin. Microbiol. 28:2139–2141. Zundel, E., J. M. Verger, M. Grayon, and R. Michel. 1992. Conjunctival vaccination of pregnant ewes and goats

CHAPTER 3.1.16 with Brucella melitensis Rev 1 vaccine: Safety and serological responses. Ann. Rech. Vet. 23:177–188. Zygmunt, M. S., H. Salih Alj Debbarh, A. Cloeckaert, and G. Dubray. 1994. Antibody response to Brucella melitensis outer membrane antigens in naturally infected and Rev1 vaccinated sheep. Vet. Microbiol. 39:33–46.

Prokaryotes (2006) 5:457–466 DOI: 10.1007/0-387-30745-1_18

CHAPTER 3.1.17 no i t cudor tnI

ot

eht

se l a i s t t ekc iR

dna

rehtO

ra l u l l ecar tnI

Introduction to the Rickettsiales and Other Intracellular Prokaryotes DAVID N. FREDRICKS

Introduction In the past, any small bacterium that required an intracellular environment for growth could be classified as belonging to the order Rickettsiales (Moulder, 1974). It is now clear that bacteria from diverse phylogenetic groups have adopted the intracellular niche, an example of convergent evolution. Most eukaryotic cells have defenses that prevent prokaryotes from invading and establishing a parasitic existence. Once these defenses are breached, however, an intracellular existence has many advantages for the suitably adapted bacterium. Within a host cell, bacteria are free from competition with most other microbes and benefit from host defenses directed against external threats. Intracellular bacteria may have access to host substrates and enzymes for metabolism. Host behaviors, such as blood feeding in insects or locomotion in vertebrate hosts, may aid the spread of intracellular bacteria. Therefore, it should not be surprising that bacteria from different phylogenetic groups have evolved to take advantage of intracellular life in eukaryotic cells. In this chapter, bacteria are organized into groups based primarily on phylogenetic inferences made from analysis of their 16S rRNA sequences (Fig. 1). Although criticized in some quarters (Pennisi, 1998), the use of 16S rRNA genes for inferring evolutionary relationships among bacteria has proven very reliable, though small subunit rRNA phylogeny may not resolve evolutionary relationships among organisms in the deepest branches of the tree of life (Woese, 2000). There is the potential for misclassification of organisms when one relies on a single gene for phylogenetic analysis. Gene insertions, deletions and transfers between organisms are possible (Nelson et al., 1999; Yap et al., 1999). However, the phylogenetic relationships suggested by analysis of 16S rRNA genes tend to be supported by phylogenetic analysis of other genes (Pace, 1997). Bacteria that form a monophyletic group with the classical rickettsiae based on phylogenetic analysis of 16S rRNA gene sequences are listed

here as belonging to the Rickettsiales. Other intracellular bacteria that have been classified traditionally as belonging to the Rickettsiales but do not cluster with this group of bacteria based on 16S rRNA analysis are listed under the Rickettsia-like prokaryotes. Figure 2 is a detailed phylogenetic tree of the bacteria described in this chapter, based on analysis of 16S rRNA. The role bacteria may play as endosymbionts of eukaryotic cells is illustrated with some examples, and the endosymbiotic hypothesis for the origin of mitochondria is discussed.

The Rickettsiales The Rickettsiales can be organized into five major genogroups based on 16S rRNA phylogeny. These groups consist of the Rickettsiae, Ehrlichiae, Neorickettsiae, Wolbachiae and Holosporae. These bacteria form a monophyletic group within the α-proteobacteria. The traditional classification scheme for the Rickettsiales is listed in Table 1, as described in Bergey’s Manual of Systematic Bacteriology (Weiss and Moulder, 1984). This chapter does not use the traditional classification scheme of families and tribes listed in Table 1 because the molecular phylogeny of these bacteria is at odds with that organizational structure.

The Rickettsia Group The rickettsiae are small (0.3–0.5 × 0.8–2.0 µm), Gram-negative, aerobic, coccobacillary organisms that are obligate intracellular parasites (Saah, 2000a). They reside free in the cytoplasm or nucleus of eukaryotic cells where they divide by binary fission and metabolize glutamate via the citric acid cycle (Weiss and Moulder, 1984). They have cell walls but no flagella. The rickettsia have been propagated in the laboratory by cocultivation with eukaryotic cells, but have not been propagated using axenic media. These bacteria frequently have a close relationship with arthropod vectors that may transmit the

se toyrakorP

458

D.N. Fredricks

CHAPTER 3.1.17

27

Thiomicrospira et al. (Coxiella, Rickettsiella) Bartonella 49

Wolbachia et al.

34

Anaplasma & Ehrlichia et al. 12

60 14

Neorickettsia et al.

Rickettsia Holospora et al. 19

92

Mitochondria

Chlamydia

0.10 Fig. 1. Phylogentic tree containing the Rickettsiales and related intracellular bacteria as inferred from 16S rRNA sequence data. This tree shows the evolutionary relationships of the major bacterial groups to each other (http://www.arb-home.de, Technical University, Munich, Germany).

Coxiella burnetii Piscirickettsia salmonis Rickettsiella grylli Bartonella henselae Wolbachia melophagi Bartonella quintana Bartonella bacilliformis Wolbachia species Wolbachia pipientis endosymbiont of Brugia malayi Ehrlichia canis Ehrlichia ewingii Ehrlichia chaffeensis Ehrlichia ruminantium "Anaplasma platys" Anaplasma equi Anaplasma phagocytophilum "Anaplasma bovis" Anaplasma marginale Neorikettsia risticii Neorickettsia sennetsu Neorickettsia helminthoeca Rickettsia rickettsii Rickettsia conorii Rickettsia akari Rickettsia typhi Rickettsia prowazekii Orientia tsutsugamushi Caedibacter caryophila Holospora obtusa Chlamydia psittaci Chlamydia pecorum Chlamydia trachomatis Chlamydia pneumoniae 0.10

Fig. 2. A detailed phylogentic tree of selected intracellular bacteria as inferred from 16S rRNA sequence data. The bar represents 0.1 nucleotide substitutions per position and is a measure of evolutionary distance (http://www.arb-home.de, Technical University, Munich, Germany).

CHAPTER 3.1.17

Introduction to the Rickettsiales and Other Intracellular Prokaryotes

459

Table 1. The traditional classification scheme for the order Rickettsiales as described in Bergey’s Manual. Order Rickettsiales

Family Rickettsiaceae

Tribe

Genera

Rickettsieae

Rickettsia Rochalimaea Coxiella Ehrlichia Cowdria Neorickettsia Wolbachia Rickettsiella Bartonella Grahamella Anaplasma Aegyptianella Haemobartonella Eperythrozoon

Ehrlicheae

Wolbachieae Bartonellaceae Anaplasmataceae

organism to mammalian hosts. The rickettsiae have very small genomes of about 1.0–1.5 million bases. The true rickettsiae can be subdivided into three major groups (spotted fever, typhus and scrub typhus groups) based on clinical characteristics of disease and phylogenetic relationships (Weisburg et al., 1989; Tamura et al., 1991). Spotted Fever Group Rickettsia rickettsii is the cause of Rocky Mountain spotted fever (RMSF) and the prototypical bacterium in the spotted fever group of rickettsiae (McDade and Newhouse, 1986). This bacterium is found in the Americas and is transmitted to humans through the bite of infected ticks. Rickettsia rickettsii infects human vascular endothelial cells, producing a vasculitis. The possibility of RMSF should be considered in patients from endemic areas presenting with fever, headache, and a rash, particularly during the summer and late spring. The rash tends to start on wrists and ankles and spreads centripetally (Walker and Raoult, 2000b). Other spotted fever group rickettsiae that produce human rickettsioses include R. conorii, R. mongolotimonae and R. slovaca (bouteonneuse fever and similar illnesses), R. akari (rickettsial pox), R. japonica (Japanese spotted fever), R. sibirica (North Asian tick typhus), R. africae (African tick bite fever), R. helvetica (perimyocarditis), R. australis (Queensland tick typhus) and R. honei (Flinders Island spotted fever; Raoult et al., 1997; Fournier et al., 2000a; Fournier et al., 2000b; Nilsson et al., 1999). The spotted fever rickettsiae have been found on every continent except Antarctica. Other spotted fever group rickettsiae, including R. montana, R. rhipicephali, R. parkeri, R. massiliae and R. aeschlimannii, have been detected in ticks, but have not been clearly linked to human disease (Walker and Raoult, 2000b).

Typhus Group Rickettsia prowazekii is the cause of epidemic or louse-borne typhus and is the prototypical bacterium from the typhus group of rickettsiae (Saah, 2000c). Like other members of the rickettsiae, R. prowazekii infects human vascular endothelial cells, producing widespread vasculitis. In contrast to RMSF, louse-borne typhus tends to occur in the winter, with the rash starting in the axillary folds and spreading centrifugally. Infection usually is transmitted from person to person by the body louse and, therefore, tends to manifest under conditions of crowding and poor hygiene. The southern flying squirrel appears to be a reservoir in the United States, but the vector involved in transmission from the flying squirrel to humans is currently unknown. The disease has a worldwide distribution. Latent human infection may occur with subsequent reactivation disease (BrillZinsser disease). Other rickettsiae in the typhus group include R. typhi and R. felis. Murine typhus is caused by transmission of R. typhi from rats, cats and opossums to humans via a flea vector (Dumler and Walker, 2000). Murine typhus is found worldwide and is endemic to areas of Texas and southern California in the United States. Rickettsia typhi infects the gut epithelium and sometimes the reproductive tissues of the flea vector. Murine typhus manifests as a nonspecific febrile illness; rash occurs in only about 50% of patients, usually as a late finding. Although R. felis is phylogenetically more closely related to the spotted fever group of rickettsiae than to the typhus group, it shares antigens with R. typhi and produces a murine typhus-like illness. Rickettsia felis has been detected in cat fleas and opossums (Higgins et al., 1996). Scrub Typhus Group Orientia tsutsugamushi is the cause of scrub typhus. Originally called Rick-

460

D.N. Fredricks

ettsia tsutsugamushi, this organism was given its own genus designation because it is phylogenetically distinct from the other rickettsiae, though closely related (Tamura et al., 1995). This bacterium has a thicker outer leaflet to its cell wall and a minimal outer slime layer, in contrast to the situation with the other genogroups of rickettsia (Weiss and Moulder, 1984). Orientia tsutsugamushi is transmitted to humans by the bite of trombiculid mites (chiggers), which are the vector and host (Saah, 2000b). Scrub typhus occurs throughout much of Asia and Australia. The clinical presentation of scrub typhus is a nonspecific febrile illness with headache and myalgias. An inoculation eschar and/or lymphadenopathy are helpful clues in its diagnosis.

The Ehrlichia Group The ehrlichia are small, Gram-negative, pleomorphic bacteria. They can be divided into several groups based on 16S rRNA phylogeny, ultrastructural features of infected cells, and pathological features of disease (Walker and Dumler, 2000a). The bacteria Ehrlichia risticii and Ehrlichia sennetsu have been designated as ehrlichiae, but it is clear from the phylogeny of these organisms that they form a distinct group with Neorickettsia helminthoeca and belong in a different but related genus (Pretzman et al., 1995; Popov et al., 1998). In this chapter, Ehrlichia risticii and Ehrlichia sennetsu are classified as neorickettsiae (next section) while awaiting further taxonomic designation. The ehrlichiae are obligate intracellular bacteria. Like chlamydiae, they exist in membrane-bound vacuoles within the cytoplasm of eukaryotic cells. Bacteria are only 0.5–1.0 µm in diameter, but can be visualized by light microscopy when they form groups of bacterial cells within vacuoles, called “morulae.” The ultrastructure of morulae formed by each genogroup of ehrlichia is distinct (Popov et al., 1998). The E. CHAFFEENSIS-E. CANIS Group The true ehrlichiae form two distinct phylogenetic clusters or genogroups of organisms. One cluster includes the agent of human monocytic ehrlichiosis (HME), E. chaffeensis, as well as E. canis, E. ewingii and E. muris. Widely found in the United States, HME is a vector-borne zoonosis transmitted by ticks of the Amblyomma and Dermacentor genus. Dogs and deer appear to be natural reservoirs of infection. Ehrlichia chaffeensis parasitizes circulating and tissue-bound macrophages. Unlike rickettsiae, ehrlichiae do not infect human endothelial cells or produce a vasculitis. The clinical presentation of HME is similar to RMSF. However, HME differs from RMSF in that a rash is found in less than half of HME

CHAPTER 3.1.17

cases, leukopenia may be present, and intracytoplasmic morulae occasionally may be detected on peripheral blood smear. Patients with HME may have multiple abnormalities evident on blood chemistry and hematology tests (Walker and Dumler, 2000a). Ehrlichia canis and E. ewingii are closely related to E. Chaffeensis. Both bacteria have been reported to cause disease in humans but are primarily canine pathogens (Buller et al., 1999). Ehrlichia canis infects macrophages in dogs, whereas E. ewingii infects granulocytes in dogs. Ehrlichia muris is another ehrlichia that clusters phylogenetically with this group, and it has been found in Japanese mice (Kawahara et al., 1993). Ticks transmit all of these bacteria. Cowdria ruminantium is an obligate intracellular bacterium that is the cause of heartwater disease, an illness of African and Caribbean ruminants. This bacterium is found within vacuoles in the cytoplasm of host endothelial cells (Moulder, 1974). Ticks of the Amblyomma genus transmit C. ruminantium. Phylogenetically, C. ruminantium falls within the E. Chaffeensis-E. Canis group of ehrlichiae and probably should be renamed Ehrlichia ruminantium (Pretzman et al., 1995). The E. PHAGOCYTOPHILA-E. EQUI Group Another genogroup of the true ehrlichiae includes the agent of human granulocytic ehrlichiosis (HGE), which appears similar or identical to E. equi and E. phagocytophila (Walker and Dumler, 2000a). The agent of HGE is transmitted to humans by ticks of the Ixodes genus and has been described in the United States and Europe. Small mammals, such as the whitefooted mouse, appear to be major reservoirs of this bacterium. Ehrlichia equi causes equine granulocytic ehrlichiosis in horses in the United States (Bullock et al., 2000). The clinical presentation of HGE is again similar to that of RMSF, except less than 10% of patients have a rash. As with HME, patients with HGE may have laboratory test abnormalities including thrombocytopenia, leukopenia and elevated serum levels of hepatic enzymes. Patients with HGE frequently have visible morulae in circulating granulocytes, whereas morulae are rarely detected in circulating monocytes of patients with HME. The phylogenetic cluster related to E. phagocytophila includes E. platys and E. bovis. Ehrlichia platys infects dogs and targets platelets and macrophages (Woody and Hoskins, 1991). The vector is unknown. Ehrlichia bovis infects cattle and buffalo. Anaplasma marginale is an obligate intracellular bacterium that infects cattle (Moulder, 1974). Anaplasmosis is found worldwide and is transmitted by ticks such as Dermacentor andersoni.

CHAPTER 3.1.17

Introduction to the Rickettsiales and Other Intracellular Prokaryotes

The bacterium occupies vacuoles within erythrocytes. Anaplasma marginale clusters phylogenetically with the E. phagocytophila-E. equi group of ehrlichiae, though it occupies a deep branch in this cluster (Drancourt and Raoult, 1994). Based on 16S rRNA phylogeny, A. marginale should be considered an ehrlichia.

The Neorickettsia Group Although related to the ehrlichiae and the rickettsiae, the neorickettsiae form a distinct phylogenetic cluster within the Rickettsiales. Ehrlichia sennetsu and E. risticii are phylogenetically very closely related organisms that do not group with the true ehrlichiae (Pretzman et al., 1995). These bacteria are closely related to Neorickettsia helminthoeca (Rikihisa, 1991). Unlike the ehrlichiae that are transmitted by tick vectors, the neorickettsiae are associated with trematode worm vectors. Neorickettsia helminthoeca is a small, pleomorphic, intracellular bacterium that causes salmon poisoning disease in canids such as coyotes and dogs (Foreyt et al., 1987). Neorickettsia helminthoeca infects cells of the canine reticuloendothelial system. A fluke vector (Nonophyetus salmincola) that infests salmonid fish transmits the disease. Dogs acquire the bacterium by eating fish containing infected flukes. Ehrlichia risticii is the cause of Potomac horse fever (PHF), a febrile colitis of horses found in North America and Europe (Palmer, 1993). It has also been associated with equine abortions. Also, PHF has been transmitted to horses by inoculation with infected trematode worms harvested from freshwater snails (Barlough et al., 1998). The natural mode of acquisition is not known, but horses may become infected by drinking or standing in trematode-infested water or by ingesting aquatic insects containing infected metacercariae. Ehrlichia risticii has been propagated in mice. There is considerable 16S rRNA sequence diversity amongst the bacteria classified as E. risticii, suggesting that there are several different strains (Wen et al., 1995). Ehrlichia sennestu is a human pathogen that has been reported to occur in Japan and Malaysia (Rapmund, 1984). It causes sennetsu fever, a mild mononucleosis-like illness marked by fever, lymphadenopathy and atypical lymphocytosis that usually resolves spontaneously. The vector has not been described, but an excellent candidate would be a trematode worm. Another bacterium related to the neorickettsiae has been detected in California rainbow trout and the trematodes infesting these fish. It is not known whether this bacterium, which has

461

not been named as yet, causes disease in mammals (Pusterla et al., 2000). An illness that was first thought to be due to E. sennetsu was later linked to another member of the neorickettsiae. People in western Japan developed an illness (Hyuga fever) after eating raw gray mullet fish (Fukuda and Yamamoto, 1981). The SF agent was isolated from the trematode worm Stellantchasmus falcatus, which infests gray mullet (Wen et al., 1996). Phylogenetic analysis of the 16S rRNA gene suggests that SF is very closely related to E. risticii.

The Wolbachia Group The Wolbachiae The Wolbachiae are small, Gram-negative, intracellular bacterial symbionts of invertebrate hosts, particularly arthropods (Dobson et al., 1999). These bacteria can infect reproductive cells and thus have the potential to be vertically transmitted by cytoplasmic inheritance. Horizontal transmission of Wolbachiae also may occur. Wolbachiae are known to affect host reproduction by inducing cytoplasmic incompatibility, parthenogenesis, male killing, feminization of genetic males, and increases in fecundity (McGraw and O’Neill, 1999; Hadfield and Axton, 1999; Hurst et al., 2000). These reproductive changes induced in host arthropods appear to promote the spread of Wolbachia endosymbionts. Wolbachia pipientis and numerous unnamed Wolbachia species make up this group (McGraw and O’Neill, 1999). Wolbachia pipientis multiplies by binary fission in host cell vacuoles. Wolbachia persica was isolated from the tick Argas persicus, now called Argas arboreus (Moulder, 1974). Phylogenetic analysis of the 16S rRNA gene shows that this bacterium does not cluster with the Wolbachiae, but rather belongs to the Francisella genus (Forsman et al., 1994). If this phylogeny proves correct, the name should probably be changed to Francisella persica. Other bacterial endosymbionts of arthropods have been described that cluster phylogenetically with the Wolbachiae, but have been named by describing the infected host. These bacteria include symbionts of Trichogramma (Genbank L02886), Bangasternus (M85266), Muscifidurax (L02882), Brugia (AF051145), Rhinocyllus (M85267), and several incompatibility symbionts (M84692, M84689, ARB [short for “arbor,” or “tree,” in Latin] database, Technical University, Munich, Germany). Wolbachia melophagi groups with the Bartonellae and is described in that section. Other Bacteria in the Wolbachia Group Bacteria that cluster with the Wolbachia based

462

D.N. Fredricks

on 16S rRNA phylogeny include bacterial symbionts associated with the algae Cosmocladium saxonicum (X79497) and Pleurastrum paucicellulare (Z47997) and a symbiont associated with the tick Haemaphysalis longcornis (AB001520). Some bacteria that are very closely related to the symbionts of Cosmocladium saxonicum have been called “Wolbachia species” (X64266) (http://www.arb-home.de, Technical University, Munich, Germany).

The Holospora Group Another group of bacteria within the Rickettsiales includes Holospora obtusa, Caedibacter caryophila, and many unnamed bacteria detected in such diverse environments as marine and rumen ecosystems. Holospora obtusa and C. caryophila are lethal symbionts of the ciliate Paramecium caudatum (Springer et al., 1993; Fujishima and Fujita, 1985). Another bacterial endosymbiont from this group has been implicated in causing necrotizing hepatopancreatitis in shrimp (Loy et al., 1996).

Rickettsia-Like Prokaryotes The Bartonellae Members of the Bartonella genus are small (0.4 × 1.5 µm), Gram-negative, aerobic rods with typical cell walls (Weiss and Moulder, 1984). They may invade erythrocytes and endothelial cells or can occupy an epicellular location. Some Bartonella have flagella. In 1993, the bacteria previously classified in the Rochalimea genus were grouped and renamed with the Bartonella genus (Breitschwerdt and Kordick, 2000). In 1995, bacteria previously in the Grahamella genus were similarly absorbed into the Bartonella genus. Bartonellae are α-proteobacteria that are phylogenetically distinct from the Rickettsiae and are related to bacteria in the Rhizobium genus. An intracellular environment is not required for growth, as these bacteria can be propagated on complex axenic media in the laboratory (Spach and Koehler, 1998). Although many of these bacteria appear to grow intracellularly, the extent of intracellular versus extracellular replication in animal host tissues is not clear. Bartonellae are important human pathogens, particularly in the immunocompromised host. A list of Bartonellae and the diseases they produce includes B. bacilliformis (Oroya fever and verruga peruana), B. henselae (cat scratch disease, bacillary angiomatosis, peliosis hepatis, bacteremia and endocarditis), B. quintana (trench fever, bacillary angiomatosis, endocarditis and bacter-

CHAPTER 3.1.17

emia), B. elizabethae (endocarditis), B. clarridgeiae (cat scratch disease), B. vinsonii arupensis (bacteremia), B. washoensis (cardiac disease) and B. grahamhii (neuroretinitis). Nonhuman mammals appear to be the primary hosts for most Bartonella species. For instance, cats are frequently infected with B. henselae, and they may have prolonged bacteremia (Heller et al., 1997; Chomel et al., 1995). Nonhuman reservoirs have not been identified for B. bacilliformis and B. quintana. Vectors for transmission of these agents are known: B. bacilliformis can be transmitted between humans via the sand fly, and B. quintana can be transmitted via the body louse. Other members of the Bartonella genus not presently associated with human disease include B. taylorii, B. doshiae, B. alsatica, B. tribocorum, B. weissii, B. koehlerae, B. vinsonii berkhoffii, B. vinsonii vinsonii, B. talpae and B. peromysci (Breitschwerdt and Kordick, 2000). A diversity of mammals and insect vectors show high rates of infection with these bacteria. Of note, the bacterium Wolbachia melophagi (Genbank X89110) does not group phylogenetically with the Wolbachiae genus, but rather groups with the Bartonellae genus. Wolbachia melophagi is closely related to B. henselae using 16S rRNA sequence phylogeny and should probably be renamed Bartonella melophagi.

The Thiomicrospira The Thiomicrospira are γ-proteobacteria that include the human and animal pathogen Coxiella burnetii, the fish pathogen Piscirickettsia salmonis, and insect symbionts, such as Rickettsiella grylli and several unnamed tick symbionts. These bacteria live within cytoplasmic vacuoles of host cells and are phylogenetically distinct from the Rickettsiae and the Bartonellae that are αproteobacteria. The Thiomicrospira are closely related to Legionella. Coxiella burnetii is a small (0.2–0.4 × 0.4– 1.0 µm), Gram-negative, obligate intracellular bacterium that passively enters cells where it replicates in the acidic environment of the phagolysosome (Maurin and Raoult, 1999). Coxiella burnetii can form an endospore that is resistant to degradation in the environment. Human infection is produced by inhalation of bacteria, though natural infection in animal populations probably involves a tick vector. In humans, C. burnetii is the cause of Q fever. A wide spectrum of organisms can harbor the bacterium, including insects, birds, fish and mammals. A bacterial pathogen of salmonid fish, Piscirickettsia salmonis, causes epizootics of disease called “piscirickettsiosis” (Mauel et al., 1999; Fryer et al., 1992). The reservoir, vector and

CHAPTER 3.1.17

Introduction to the Rickettsiales and Other Intracellular Prokaryotes

mode of acquisition for this bacterium are not known. Rickettsiella grylli is another member of the Thiomicrospira and is not related to the Rickettsiae despite the association implied by its name (Roux et al., 1997; Frutos et al., 1994). Rickettsiella grylli has a cricket host. The bacterium enters a host cell vacuole where it divides. Like chlamydia, these bacteria undergo a developmental cycle consisting of stages, including a large particle that replicates and a small dense particle that is infectious. The bacteria may form disk shapes and condense into crystalline bodies. Other members of this taxonomic group include symbionts of Rhipicephalus and Ornithodoros ticks.

The Chlamydiae Chlamydiae are small bacteria (0.2–1.5 µm) that replicate within the phagosome of eukaryotic cells. Whereas rickettsiae exit the phagosome after phagocytosis and multiply in the cell cytoplasm, the chlamydiae (and ehrlichiae) remain in the phagosome and prevent fusion with lysosomes (Weiss and Moulder, 1984). Chlamydia do not metabolize glutamate, in contrast to the rickettsiae. Chlamydiae have some structural features of Gram-negative bacteria, but their cell wall lacks peptidoglycan. Chlamydiae have a biphasic life cycle. The infectious stage is more compact and called “the elementary body,” whereas the replicative stage is larger and called “the reticulate body.” The chlamydiae form their own distinct phylogenetic group within the eubacteria (Everett, 2000) and are peripherally related to the planctomycetes, a group of aquatic and terrestrial bacteria that also lack peptidoglycan in their cell walls (Weisburg et al., 1986). The currently defined species of chlamydia include C. psittaci, C. pecorum, C. trachomatis and C. pneumoniae. Chlamydia pecorum is exclusively an animal pathogen, whereas the other three chlamydiae can cause human disease. Chlamydia psittici causes disease in birds and mammals. Humans acquire the bacterium via inhalation of aerosols generated from infected animals, with resulting pneumonia (Vanrompay et al., 1995). Closely related to C. psittici, C. pecorum causes disease in animals such as ruminants, swine and marsupials (Fukushi and Hirai, 1993). In humans, C. trachomatis causes ocular (trachoma), genital disease (lymphogranuloma venereum) and, less commonly, other syndromes (Stamm, 1999). Chlamydia pneumoniae, exclusively a human pathogen producing respiratory disease (Kuo et al., 1995), has also been linked to atherosclerosis in humans.

463

Prokaryotic Endosymbionts of Eukaryotic Cells The relationships between prokaryotic and eukaryotic cells are discussed in detail (see section on Symbiotic Associations in this Chapter). This section will give examples of how some intracellular prokaryotes have evolved with their eukaryotic hosts. Wolbachia pipientis, a member of the Rickettsiales, is a common cytoplasmic symbiont of insects. This bacterium is widely distributed in host tissues and can be transmitted vertically in the cytoplasm of infected host eggs, though horizontal transmission may also occur. The numerous effects of W. pipientis on insect reproduction increase the number of female offspring while suppressing the number of male offspring (McGraw and O’Neill, 1999; Stouthamer et al., 1999). Because female insects can vertically transmit W. pipientis to their offspring, this strategy benefits the endosymbiont. One mechanism for producing this result is crossing incompatibility. Infected females can mate with infected or uninfected males, but uninfected females cannot mate with infected males. Another mechanism promoting female representation is the direct killing of male embryos by W. pipientis. Buchnera aphidicola is an obligate intracellular symbiont of aphids that is vertically transmitted and colonizes specialized host cells called “bacteriocytes” (Clark et al., 2000). This bacterium is not a member of the Rickettsiales, but rather is a member of the γ-proteobacteria. The complete genome of Buchnera has been sequenced (Shigenobu et al., 2000). Buchnera has genes for the biosynthesis of essential amino acids required by the host, but lacks those for nonessential amino acids produced by the host, demonstrating complete biochemical interdependence. Indeed, eradication of bacteria from bacteriocytes is lethal for aphids.

Mitochondria: The Ultimate Prokaryotic Endosymbiont Mitochondria are specialized aerobic energy producing organelles of eukaryotic cells. Like chloroplasts, mitochondria have unique genomes that are distinct from the nuclear genomes of their associated eukaryotic cells (Gray, 1993). When mitochondrial DNA from diverse eukaryotes are compared, there are striking differences in the size, organization and gene content of these genomes (Gray et al., 1999). Yet, phylogenetic analyses suggest that all mitochondrial genomes descended from a common ancestor related to the α-proteobacteria (Lang et al.,

464

D.N. Fredricks

1999). The most complete mitochondrial genome has been found in the flagellated protozoan Reclinomonas americana (Lang et al., 1997). This genome contains 97 genes, including genes for bacterial rRNA and genes encoding a eubacteria-like multicomponent RNA polymerase. Of the bacterial genomes that have been sequenced, that of Rickettsia prowazekii is most closely related to the mitochondrial genome (Andersson et al., 1998). These data suggest a bacterial origin for mitochondria. According to the endosymbiont hypothesis (Gray and Doolittle, 1982), about 1.5–2.0 billion years ago an α-proteobacterium entered a proto-eukaryotic cell where it established an endosymbiotic relationship. Progressive gene loss and specialization of function led to the development of mitochondria as known today. Mitochondria allowed eukaryotic cells to generate energy without using the plasma membrane. The host cell provided substrates for energy production and a protected niche for replication. Regulation of the mitochondrial genome eventually shifted to nuclear control. The hydrogen hypothesis is an extension of the endosymbiont hypothesis for the origin of mitochondria (Rotte et al., 2000; Bui et al., 1996). This theory holds that a hydrogen-requiring archaebacterium engulfed a hydrogen-producing αproteobacterium. Specialization of function by the endosymbiont led to the formation of hydrogenosomes for anaerobic energy production in eukaryotic cells. Hydrogenosomes occur in early amitochondric eukaryotic cells, such as Trichomonads. Although lacking in DNA, hydrogenosomes have proteins (e.g., heat shock proteins) that are phylogenetically related to mitochondrial proteins (Bui et al., 1996; Dyall et al., 2000). These data suggest a common origin for hydrogenosomes and mitochondria, though it is not clear if there was a single endosymbiotic event leading to two different organelles or multiple introductions with closely related prokaryotes. Mitochondria appear to have originated from the introduction of a single α-proteobacterial endosymbiont into a eukaryotic or protoeukaryotic cell (Gray et al., 1999). Although theoretically possible, a model in which different endosymbiont bacteria formed mitochondria is not supported by the data. Mitochondria are most closely related to the rickettsiae. However, one cannot conclude that the mitochondrial ancestor also belonged to this genus. It is possible that an α-proteobacterial ancestor gave rise to both the rickettsiae and the mitochondrial endosymbiont. The rickettsial and mitochondrial genomes are quite small. Both genomes are likely to have sustained gene loss over evolutionary time related to their common intracellular

CHAPTER 3.1.17

niche. Thus, the extensive loss of mitochondrial genes does not necessarily imply a rickettsial origin. If evolutionary success is measured by the ability to pass a genome to succeeding generations, then the mitochondrial endosymbiont has been fantastically successful. Mitochondria in eukaryotic cells provide evidence that the strategy of intracellular prokaryotic existence is ancient and durable.

Literature Cited Andersson, S. G., A. Zomorodipour, J. O. Andersson, et al. 1998. The genome sequence of Rickettsia prowazekii and the origin of mitochondria. Nature 396:133–140. Barlough, J. E., G. H. Reubel, J. E. Madigan, L. K. Vredevoe, P. E. Miller, and Y. Rikihisa. 1998. Detection of Ehrlichia risticii, the agent of Potomac horse fever, in freshwater stream snails (Pleuroceridae: Juga spp.) from northern California. Appl. Environ. Microbiol. 64:2888–2893. Breitschwerdt, E. B., and D. L. Kordick. 2000. Bartonella infection in animals: Carriership, reservoir potential, pathogenicity, and zoonotic potential for human infection. Clin. Microbiol. Rev. 13:428–438. Bui, E. T., P. J. Bradley, and P. J. Johnson. 1996. A common evolutionary origin for mitochondria and hydrogenosomes. Proc. Natl. Acad. Sci. USA 93:9651–9656. Buller, R. S., M. Arens, S. P. Hmiel, et al. 1999. Ehrlichia ewingii, a newly recognized agent of human ehrlichiosis. N. Engl. J. Med. 341:148–155. Bullock, P. M., T. R. Ames, R. A. Robinson, B. Greig, M. A. Mellencamp, and J. S. Dumler. 2000. Ehrlichia equi infection of horses from Minnesota and Wisconsin: Detection of seroconversion and acute disease investigation. J. Vet. Intern. Med. 14:252–257. Chomel, B. B., R. C. Abbott, and R. W. Kasten, et al. 1995. Bartonella henselae prevalence in domestic cats in California: Risk factors and association between bacteremia and antibody titers. J. Clin. Microbiol. 33:2445–50. Clark, M. A., N. A. Moran, P. Baumann, and J. J. Wernegreen. 2000. Cospeciation between bacterial endosymbionts (Buchnera) and a recent radiation of aphids (Uroleucon) and pitfalls of testing for phylogenetic congruence. Evolution Int J Org Evolution 54:517–525. Dobson, S. L., K. Bourtzis, H. R. Braig, et al. 1999. Wolbachia infections are distributed throughout insect somatic and germ line tissues. Insect Biochem. Molec. Biol. 29:153– 160. Drancourt, M., and D. Raoult. 1994. Taxonomic position of the rickettsiae: Current knowledge. FEMS Microbiol. Rev. 1:13–24. Dumler, J. S., and D. H. Walker. 2000. Rickettsia typhi (Murine typhus). In: G. L. Mandell, J. E. Bennett, and R. Dolin (Eds.) Principles and Practice of Infectious Diseases. Churchill Livingstone. Philadelphia, PA. 2:2053–2056. Dyall, S. D., C. M. Koehler, M. G. Delgadillo-Correa, et al. 2000. Presence of a member of the mitochondrial carrier family in hydrogenosomes: Cnservation of membranetargeting pathways between hydrogenosomes and mitochondria. Molec. Cell. Biol. 20:2488–97.

CHAPTER 3.1.17

Introduction to the Rickettsiales and Other Intracellular Prokaryotes

Everett, K. D. 2000. Chlamydia and Chlamydiales: More than meets the eye. Vet. Microbiol. 75:109–126. Foreyt, W. J., J. R. Gorham, J. S. Green, C. W. Leathers, and B. R. LeaMaster. 1987. Salmon poisoning disease in juvenile coyotes: Clinical evaluation and infectivity of metacercariae and rickettsiae. J. Wildl. Dis. 23:412– 417. Forsman, M., G. Sandstrom, and A. Sjostedt. 1994. Analysis of 16S ribosomal DNA sequences of Francisella strains and utilization for determination of the phylogeny of the genus and for identification of strains by PCR. Int. J. Syst. Bacteriol. 44:38–46. Fournier, P. E., F. Grunnenberger, B. Jaulhac, G. Gastinger, and D. Raoult. 2000a. Evidence of Rickettsia helvetica infection in humans, eastern France. Emerg. Infect. Dis. 6:389–392. Fournier, P. E., H. Tissot-Dupont, H. Gallais, and D. R. Raoult. 2000b. Rickettsia mongolotimonae: A rare pathogen in France. Emerg. Infect. Dis. 6:290–292. Frutos, R., B. A. Federici, B. Revet, and M. Bergoin. 1994. Taxonomic studies of Rickettsiella, Rickettsia, and Chlamydia using genomic DNA. J. Invertebr. Pathol. 63:294–300. Fryer, J. L.,C. N. Lannan, S. J. Giovannoni, and N. D. Wood. 1992. Piscirickettsia salmonis gen. nov., sp. nov., the causative agent of an epizootic disease in salmonid fishes. Int. J. Syst. Bacteriol. 42:120–126. Fujishima, M., and M. Fujita. 1985. Infection and maintenance of Holospora obtusa, a macronucleus-specific bacterium of the ciliate Paramecium caudatum. J. Cell. Sci. 76:179–187. Fukuda, T., and S. Yamamoto. 1981. Neorickettsia-like organism isolated from metacercaria of a fluke, Stellantchasmus falcatus. Japan. J. Med. Sci. Biol. 34:103–107. Fukushi, H., and K. Hirai. 1993. Chlamydia pecorum— the fourth species of genus Chlamydia. Microbiol. Immunol. 37:516–522. Gray, M. W., and W. F. Doolittle. 1982. Has the endosymbiont hypothesis been proven? Microbiol. Rev. 46:1–42. Gray, M. W. 1993. Origin and evolution of organelle genomes. Curr. Opin. Genet. Dev. 3:884–890. Gray, M. W., G. Burger, and B. F. Lang. 1999. Mitochondrial evolution. Science 283:1476–1481. Hadfield, S. J., and J. M. Axton. 1999. Germ cells colonized by endosymbiotic bacteria. Nature 402:482. Heller, R., M. Artois, V. Xemar, et al. 1997. Prevalence of Bartonella henselae and Bartonella clarridgeiae in stray cats. J. Clin. Microbiol. 35:1327–1331. Higgins, J. A., S. Radulovic, M. E. Schriefer, and A. F. Azad. 1996. Rickettsia felis: A new species of pathogenic rickettsia isolated from cat fleas. J. Clin. Microbiol. 34:671– 674. Hurst, G. D., A. P. Johnson, D., von der Schulenburg, J. H., and Y. Fuyama. 2000. Male-killing Wolbachia in Drosophila: A temperature-sensitive trait with a threshold bacterial density. Genetics 156:699–709. Kawahara, M., C. Suto, Y. Rikihisa, S. Yamamoto, and Y. Tsuboi. 1993. Characterization of ehrlichial organisms isolated from a wild mouse. J. Clin. Microbiol. 31:89– 96. Kuo, C. C., L. A. Jackson, L. A. Campbell, and J. T. Grayston. 1995. Chlamydia pneumoniae (TWAR). Clin. Microbiol. Rev. 8:451–461. Lang, B. F., G. Burger, C. J. O’Kelly, et al. 1997. An ancestral mitochondrial DNA resembling a eubacterial genome in miniature. Nature 387:493–497.

465

Lang, B. F., E. Seif, M. W. Gray, C. J. O’Kelly, and G. Burger. 1999. A comparative genomics approach to the evolution of eukaryotes and their mitochondria. J. Eukaryot. Microbiol. 46:320–326. Loy, J, K., F. E. Dewhirst, W. Weber, et al. 1996. Molecular phylogeny and in situ detection of the etiologic agent of necrotizing hepatopancreatitis in shrimp. Appl. Environ. Microbiol. 62:3439–3445. Mauel, M. J., S. J. Giovannoni, and J. L. Fryer. 1999. Phylogenetic analysis of Piscirickettsia salmonis by 16S, internal transcribed spacer (ITS) and 23S ribosomal DNA sequencing. Dis. Aquat. Organ. 35:115–123. Maurin, M., and D. Raoult. 1999. Q fever. Clin. Microbiol. Rev. 12:518–553. McDade, J. E., and V. F. Newhouse. 1986. Natural history of Rickettsia rickettsii. Ann. Rev. Microbiol. 40:287–309. McGraw, E. A., and S. L. O’Neill. 1999. Evolution of Wolbachia pipientis transmission dynamics in insects. Trends Microbiol. 7:297–302. Moulder, J. W. 1974. Rickettsiales. In: R. E. Buchanan and N. E. Gibbons (Eds.) Bergey’s Manual of Determinative Bacteriology. Waverly Press. Baltimore, MD. 882–915. Nelson, K. E., R. A. Clayton, S. R. Gill, et al. 1999. Evidence for lateral gene transfer between Archaea and bacteria from genome sequence of Thermotoga maritima. Nature 399:323–329. Nilsson, K., O. Lindquist, and C. Pahlson. 1999. Association of Rickettsia helvetica with chronic perimyocarditis in sudden cardiac death. Lancet 354:1169–1173. Pace, N. R. 1997. A molecular view of microbial diversity and the biosphere. Science 276:734–740. Palmer, J. E. 1993. Potomac horse fever. Vet. Clin. North Am. Equine Pract. 9:399–410. Pennisi, E. 1998. Geome data shake Tree of Life. Science 280:672–674. Popov, V. L., V. C. Han, S. M. Chen, et al. 1998. Ultrastructural differentiation of the genogroups in the genus Ehrlichia. J. Med. Microbiol. 47:235–251. Pretzman, C., D. Ralph, D. R. Stothard, P. A. Fuerst, and Y. Rikihisa. 1995. 16S rRNA gene sequence of Neorickettsia helminthoeca and its phylogenetic alignment with members of the genus Ehrlichia. Int. J. Syst. Bacteriol. 45:207–211. Pusterla, N., E. Johnson, J. Chae, et al. 2000. Molecular detection of an ehrlichia-like agent in rainbow trout (Oncorhynchus mykiss) from northern California. Vet. Parasitol. 92:199–207. Raoult, D., P. Berbis, V. Roux, W. Xu, and M. Maurin. 1997. A new tick-transmitted disease due to Rickettsia slovaca [letter]. Lancet 350:112–113. Rapmund, G. 1984. Rickettsial diseases of the Far East: New perspectives. J. Infect. Dis. 149:330–338. Rikihisa, Y. 1991. Cross-reacting antigens between Neorickettsia helminthoeca and Ehrlichia species, shown by immunofluorescence and Western immunoblotting. J. Clin. Microbiol. 29:2024–2029. Rotte, C., K. Henze, M. Muller, and W. Martin. 2000. Origins of hydrogenosomes and mitochondria. commentary. Curr. Opin. Microbiol. 3:481–486. Roux, V., M. Bergoin, N. Lamaze, and D. Raoult. 1997. Reassessment of the taxonomic position of Rickettsiella grylli. Int. J. Syst. Bacteriol. 47:1255–1257. Saah, A. J. 2000a. Introduction to Rickettsioses and Ehrlichioses. In: G. L. Mandell, J. E. Bennett, and R. Dolin (Eds.) Principles and Practice of Infectious

466

D.N. Fredricks

Diseases. Churchill Livingstone. Philadelphia, PA. 2:2033–2035. Saah, A. J. 2000b. Orientia tsutsugamushi (Scrub typhus). In: G. L. Mandell, J. E. Bennett, and R. Dolin (Eds.) Principles and Practice of Infectious Diseases. Churchill Livingstone. Philadelphia, PA. 2:2056–2057. Saah, A. J. 2000c. Rickettsia prowazekii (Epidemic or louseborne typhus). In: G. L. Mandell, J. E. Bennett, and R. Dolin (Eds.) Principles and Practice of Infectious Diseases. Churchill Livingstone. Philadelphia, PA. 2:2050–2053. Shigenobu, S., H. Watanabe, M. Hattori, Y. Sakaki, and H. Ishikawa. 2000. Genome sequence of the endocellular bacterial symbiont of aphids Buchnera sp. APS. Nature 407:81–86. Spach, D. H., and J. E. Koehler. 1998. Bartonella-associated infections. Infect. Dis. Clin. North Am. 12:137–155. Springer, N., W. Ludwig, R. Amann, H. J. Schmidt, H. D. Gortz, and K. H. Schleifer. 1993. Occurrence of fragmented 16S rRNA in an obligate bacterial endosymbiont of Paramecium caudatum. Proc. Natl. Acad. Sci. USA 90:9892–9895. Stamm, W. E. 1999. Chlamydia trachomatis infections: Progress and problems. J. Infect. Dis. 179, Suppl. 2:S380– S383. Stouthamer, R., J. A. Breeuwer, and G. D. Hurst. 1999. Wolbachia pipientis: Microbial manipulator of arthropod reproduction. Ann. Rev. Microbiol. 53:71–102. Tamura, A., H. Urakami, and N. Ohashi. 1991. A comparative view of Rickettsia tsutsugamushi and the other groups of rickettsiae. Eur. J. Epidemiol. 7:259–269. Tamura, A., N. Ohashi, H. Urakami, and S Miyamura. 1995. Classification of Rickettsia tsutsugamushi in a new genus, Orientia gen. nov., as Orientia tsutsugamushi comb. nov. Int. J. Syst. Bacteriol. 45:589–591. Vanrompay, D., R. Ducatelle, and F. Haesebrouck. 1995. Chlamydia psittaci infections: A review with emphasis on avian chlamydiosis. Vet. Microbiol. 45:93–119.

CHAPTER 3.1.17 Walker, D. H., and J. S. Dumler. 2000a. Ehrlichia chaffeensis, Ehrlichia phagocytophila, and other Ehrlichia. In: G. L. Mandell, J. E. Bennett, and R. Dolin (Eds.) Principles and Practice of Infectious Diseases. Churchill Livingstone. Philadelphia, PA. 2:2057–2065. Walker, D. H., and D. Raoult. 2000b. Rickettsia rickettsii and other spotted fever group Rickettsiae. In: G. L. Mandell, J. E. Bennett, and R. Dolin (Eds.) Principles and Practice of Infectious Diseases. Churchill Livingstone. Philadelphia, PA. 2:2035–2042. Weisburg, W. G., T. P. Hatch, and C. R. Woese. 1986. Eubacterial origin of chlamydiae. J. Bacteriol. 167:570–574. Weisburg, W. G., M. E. Dobson, J. E. Samuel, et al. 1989. Phylogenetic diversity of the Rickettsiae. J. Bacteriol. 171:4202–4206. Weiss, E., and J. W. Moulder. 1984. Rickettsias and Chlamydias. In: N. R. Kreig and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 687–739. Wen, B., Y. Rikihisa, and P. A. Fuerst. 1995. Chaichanasiriwithaya W: Diversity of 16S rRNA genes of new Ehrlichia strains isolated from horses with clinical signs of Potomac horse fever. Int. J. Syst. Bacteriol. 45:315–318. Wen, B., Y. Rikihisa, S. Yamamoto, N. Kawabata, and P. A. Fuerst. 1996. Characterization of the SF agent, an Ehrlichia sp. isolated from the fluke Stellantchasmus falcatus, by 16S rRNA base sequence, serological, and morphological analyses. Int. J. Syst. Bacteriol. 46:149– 154. Woese, C. R. 2000. Interpreting the universal phylogenetic tree. Proc. Natl. Acad. Sci. USA 97:8392–8396. Woody, B. J., and J. D. Hoskins. 1991. Ehrlichial diseases of dogs. Vet. Clin. North Am. Small Anim. Pract. 21:75–98. Yap, W. H., Z. Zhang, and Y. Wang. 1999. Distinct types of rRNA operons exist in the genome of the actinomycete Thermomonospora chromogena and evidence for horizontal transfer of an entire rRNA operon. J. Bacteriol. 181:5201–5209.

Prokaryotes (2006) 5:467–492 DOI: 10.1007/0-387-30745-1_19

CHAPTER 3.1.18 ehT

suneG

a l l enot raB

The Genus Bartonella MICHAEL F. MINNICK AND BURT E. ANDERSON

Phylogeny Bartonella species share many general characteristics. Members of the genus are small (approximately 0.3 µm × 1 µm), Gram-negative, pleomorphic coccobacilli. The bacteria are facultative intracellular pathogens, many of which employ hemotrophy (infection of erythrocytes) as a parasitic strategy. All members of the genus are notoriously fastidious and grow slowly in vitro. Bartonella species have been shown to infect a variety of mammalian hosts, and at least three species (B. bacilliformis, B. henselae and B. quintana) are relatively common human pathogens. Vector-mediated transmission is another common theme within the genus. Bartonella spp. are typically transmitted between mammalian hosts by arthropods, with each bacterial species transmitted by a particular insect vector. Reservoirs for Bartonella spp. include a diverse array of mammals. Based on 16S rRNA gene sequence comparisons, members of the genus Bartonella fall into the α-2 subgroup of the Proteobacteria (O’Connor et al., 1991). The genera closest to Bartonella include Brucella, Rhizobium and Agrobacterium. In general, the current genus Bartonella is relatively homogeneous with members exhibiting greater than 95% identity among aligned 16S rRNA gene sequences, resulting in a somewhat tightly branched phylogenetic tree (Fig. 1). Phylogenetic relationships determined using other loci, including gltA (the citrate synthase gene; Birtles and Raoult, 1996), ftsZ (a gene specifying a cell division [tubulin-like] protein; Kelly et al., 1998; Ehrenborg et al., 2000), and the 17-kDa antigen gene (Sweger et al., 2000), have resulted in dendrograms similar to that derived from the 16S rRNA gene sequences.

Taxonomy The genus Bartonella has recently undergone a major taxonomic reorganization. Since the description of Bartonella bacilliformis in 1909, this species was the only representative from the

genus described by A. L. Barton (Barton, 1909). However, bacteria once constituting the genera Rochalimaea and Grahamella were reclassified into the genus Bartonella (Brenner et al., 1993; Birtles et al., 1995), increasing the number of potential species to 15. Three Bartonella species are etiologic agents of major emerging infectious diseases in humans (Table 1). An additional four species (and two subspecies) of Bartonella also have been associated with human disease or described as human pathogens in isolated case reports (Table 2). Finally, ten species have not been associated with human disease but have been found in a variety of animal hosts (Table 3). The group’s potential impact on human health and their ubiquitous presence in animal reservoirs have fueled an interest in understanding the basic biology of these bacteria as well as the epidemiology, clinical presentation and natural history of the resultant diseases. The Bartonellaceae family has been removed from the order Rickettsiales; a group that originally included the Anaplasmataceae, Rickettsiaceae and Bartonellaceae. Members of the family Bartonellaceae share some rickettsia-like characteristics such as small size (e.g., the “virus-like particles” of B. bacilliformis), its transmission to humans through an arthropod vector (sandflies), and its ability to live within host cells (Moulder, 1974). However, they are not true obligate intracellular parasites like the rickettsiae, and several phenotypic properties differ between members of Bartonellaceae and the rickettsiae. For instance, Bartonella, Grahamella and Rochalimaea species and a few species of Wolbachia were the only Rickettsiales members that can be cultured in vitro, whereas true rickettsiae are strictly obligate intracellular parasites of eukaryotic cells. In addition, B. bacilliformis was the only flagellated member within the order (Weinman, 1974). Recently obtained DNA relatedness data, mainly 16S rRNA gene sequences, have clearly shown that Rochalimaea and Bartonella are more closely related to each other than to any rickettsia and most closely related to members of the α-2 subgroup of the Proteobacteria

468

M.F. Minnick and B.E. Anderson

CHAPTER 3.1.18

Table 1. The three major human pathogens in the genus Bartonella. Species bacilliformis henselae quintana

Common manifestation(s)

Vector(s)

Oroya fever, verruga peruana Cat-scratch disease, endocarditis, bacillary angiomatosis, bacillary peliosis, bacteremic syndrome Trench fever, endocarditis, bacillary angiomatosis, bacteremic syndrome

Reservoir(s)

Sandflies Cats, fleas

Humans Cats

Body louse

Humans

Table 2. Bartonella species associated with human disease.a Species clarridgeiae elizabethae grahamii vinsonii sub sp. aurepensis vinsonii sub sp. berkhoffii

Clinical manifestation

Vector

Reservoir(s)

Reference

Cat-scratch disease? Endocarditis Retinitis Bacteremia Endocarditis

Cats Unknown Unknown Ticks? Unknown

Cats Unknown Rodents Domestic dog, rodents Unknown

Kordick et al., 1997 Daly et al., 1993 Kerkhoff et al., 1999 Welch et al., 1999 Roux et al., 2000

a

Includes species associated with disease through serology or single case reports of disease associated with isolation or detection.

B. clarridgeiae B. vinsonii berkhoffii B. vinsonii arupensis B. grahamii B. tribocorum B. elizabethae B. washoensis B. quintana B. koehlerae B. weissii B. henselae B. alsatica B. vinsonii vinsonii B. taylorii B. doshiae B. bacilliformis Brucalla abortus 0.01 Fig. 1. Phylogenetic tree showing relationships of Bartonella spp. The 16S rRNA gene was aligned with over 1,272 bases. Brucella abortus is shown as an outgroup. Bar indicates a 1% nucleotide divergence. (From Breitschwerdt and Kordick, 2000, with permission.)

class, especially the Rhizobiaceae (Birtles et al., 1991; O’Connor et al., 1991; Brenner et al., 1991; Relman et al., 1992). On the basis of these findings, Rochalimaea species were subsequently reclassified as Bartonella, and the Bartonellaceae family (containing both the Grahamella and Bartonella genera) was removed from the Rickettsiales order (Brenner et al., 1993). A subsequent study reclassified five Grahamella species as bartonellae based upon DNA relatedness data and phenotypic characteristics, thereby eliminating the Grahamella genus (Birtles et al., 1995). The Bartonella genus presently contains the 18 species as listed in Tables 1–3. The phylogenetic relationship of Bartonella to other α-Proteobacteria indicates that the closest relatives to Bartonella include Agrobacterium, Rhizobium and

Table 3. Bartonella species that have not been demonstrated to be pathogenic for humans. Species alsatica doshiae grahamii koehlerae peromysci talpae taylorii tribocorum vinsonii sub sp. vinsonii washoensis weisii

Host(s)

Reference(s)

Rabbits Rodents Rodents, insectivores Domestic cat Rodents Moles Rodents Rats Voles

Heller et al., 1999 Birtles et al., 1995 Birtles et al., 1995 Droz et al., 1999 Birtles et al., 1995 Birtles et al., 1995 Birtles et al., 1995 Heller et al., 1998 Weiss and Dasch, 1982 Unpublished Unpublished

Ground squirrels Cats

Brucella species, the latter being the most closely related (Brenner et al., 1993). At first glance this grouping seems diverse, but its members do show similar natural histories. Specifically, all four genera have evolved towards a parasitic or mutualistic association with eukaryotic cells. Bartonella spp. and Brucella spp. are both intracellular parasites of mammalian cells, whereas Agrobacterium and Rhizobium species can parasitize or mutualistically associate with plant cells, respectively. None of these bacteria are obligate parasites, and all can be cultivated in vitro. Interestingly, human diseases caused by Brucella and Agrobacterium spp. share superficial similarities with bartonelloses. Like with Bartonella spp., opportunistic infections with Agrobacterium radiobacter mainly occur in immunocompromised patients and can cause bacteremia and endocarditis (Edmond et al., 1993; Freney et al., 1985; Plotkin, 1980; Southern, 1996). In addition, some manifestations of brucellosis resemble the symptoms associated with

CHAPTER 3.1.18

infection by B. henselae or B. quintana. For instance, brucellosis is a febrile illness characterized by granulomatous tissue in the lymph nodes, liver, spleen and bone and is accompanied by lymphadenopathy, bacteremia and occasionally endocarditis (Wilfert, 1992).

Habitat Bartonella species have been isolated or detected in a wide range of animal species. Included in the list of potential animal reservoirs are cats, dogs, rodents, rabbits and cattle as well as a diverse group of wild animals including wildcats (bobcats, pumas and mountain lions), coyotes, deer, elk and foxes (for review, see Breitschwerdt, 2000). At least three species have been identified as major human pathogens (Table 1), and their disease syndromes are described below. An additional four species, including two subspecies, have been associated with human disease either indirectly or through isolated case reports (Table 2). However, their identification as major human pathogens awaits further reports associating them with human disease. Finally, currently eleven other species of Bartonella have only been isolated from animals (Table 3). It should be stressed that separation of individual species into human and animal agents is likely to change as specific diagnostic tests for all Bartonella species are developed and applied to testing human samples. Transmission of Bartonella to humans occurs through an insect vector for most Bartonella species. The list of vectors associated with transmission includes flies, fleas, ticks, lice and mites (Tables 1 and 2). The possibility of direct mechanical transmission of B. henselae from cats to humans resulting in cat-scratch disease has been suggested. The role of the cat flea in this process seems likely and may involve the contamination of cat claws with infected flea feces. Transmission of B. henselae among cats appears to require the cat flea as a vector (Chomel et al., 1996). The epidemiological association of cat fleas with cases of human cat-scratch disease further supports the idea that contaminated flea feces is required for transmission (Zangwill et al., 1993).

Isolation Isolation of Bartonella spp. from clinical specimens requires extended incubation times and specialized media. In addition, direct plating of specimens prepared by the lysis-centrifugation method (Wampole Laboratories, Cranbury, NJ) sometimes enhances isolation from blood

The Genus Bartonella

469

(Slater et al., 1990). Alternatively, homogenized tissue can serve as the inoculum on blood plates or endothelial cell monolayers (Koehler et al., 1992). Blood or chocolate agar plates with a complex base medium (trypticase soy, heart infusion or Columbia agar) are used for culturing. The pH of the medium should be 7.0 to 7.5 for optimal growth. Bartonella spp. also require high humidity and, with the exception of B. bacilliformis, prefer 5% CO2 within the growth chamber. The slow growth rate of these bacteria can compound primary isolation and often requires incubation times exceeding 3–4 weeks to visualize colonies. The combination of a slow growth rate and fastidious nature can make isolation and axenic culture of Bartonella spp. a challenging undertaking. In fact, endocarditis isolates of B. henselae were only recently obtained (Drancourt et al., 1996). Following isolation and pure culture of a strain, identification of the suspected Bartonella species can be accomplished using the techniques described below in “Identification.”

Identification In addition to a number of growth characteristics, structural, biochemical and genetic properties have been used to assist in the identification of Bartonella isolates. In general, polymerase chain reaction (PCR) amplification coupled with sequencing, restriction fragment-length polymorphism (RFLP) analysis or genetic probing recently has gained favor for rapid and definitive identification of isolates.

Colony Morphology Bartonella colonies, typically small (1–3 mm diameter) and round, range from transluscent to opaque (white or cream) in color. Colonies may display a dry-to-mucoid phase variation with repeated passage. Colonies obtained from lowpassage isolates typically adhere to and pit the medium, but this phenotype disappears following repeated passage in vitro. Colonies of B. bacilliformis freely interconvert between a small, transluscent round colony (T1) to a larger colony with an irregular edge (T2; Walker and Winkler, 1981). Colonies from primary isolates can take up to four weeks before becoming visible; however, growth is much more rapid (2–5 days) in subsequent passages. It has been suggested that this colony variation may correlate with antigenic phase variation seen with B. henselae and B. quintana, with fresh isolates being more highly reactive with human sera than isolates that have undergone extensive laboratory passaging (Regnery and Tappero, 1995).

470

M.F. Minnick and B.E. Anderson

CHAPTER 3.1.18

Cell Morphology Bartonella cells are Gram negative, non-acid fast, pleomorphic rods. Bartonella spp. stain poorly with safranin, but stain well with Giemsa or Gimenez stains. Cells are typically coccobacilli or slightly curved rods with a polar enlargement(s). Cells also can be coccoid, beaded, filamentous or in chains. Annular (ring forms) and autoaggregates also can occur. Cell size is uniformly less than 3 µm in the greatest dimension, with most cells measuring 0.5 µm × 1.0 µm. Figure 2 shows examples of the two basic cell morphologies observed in Bartonella spp. including a flagellated and nonflagellated cell.

therefore test results must be judged cautiously. If hemin is added to physiological test media, test results for acid production from carbohydrates (lactose, maltose and saccharose), hippurate hydrolysis, pyrazinamidase and Voges-Proskauer can be used to differentiate B. henselae from B. quintana (Drancourt and Raoult, 1993). RapID ANA panels (Innovative Diagnostic Systems, Inc., Norcross, GA; Daly et al., 1993; Clarridge et al., 1995), DNA hybridization (Regnery et al., 1991; Welch et al., 1992) and pre-formed peptidases (Welch et al., 1992) also have been used with varying success for the identification of Bartonella species.

Polymerase Chain Reaction

Biochemical Tests Bartonella spp. are nonfermentative aerobes with unremarkable physiology. For this reason biochemical tests are usually not conclusive for species identification. One potential problem with standard biochemical tests is that they do not include hemin for Bartonella growth, and

A

B

The PCR is a sensitive and specific tool for identifying “non-culturable” Bartonella spp. in human samples or for confirming presumptive identification of isolates. Frequently, PCR has been used to amplify portions, or all, of the 16S rDNA gene from suspected Bartonella spp. The 16S rDNA PCR product is subsequently sequenced, and the data compared to known 16S rDNA sequences in GenBank to confirm presumptive identification (Relman et al., 1990; Regnery et al., 1992a; Koehler et al., 1992; Hadfield et al., 1993). To streamline identification, PCR strategies also have been devised using Bartonella genus-specific or species-specific targets. Targets have included the 16S rDNA gene (Dauga et al., 1996), a heat shock protein or stress endopeptidase gene (htrA; Anderson et al., 1993), the gltA gene (Regnery et al., 1991; Birtles and Raoult, 1996; Patel et al., 1999), the ribC gene (Bereswill et al., 1999), the ftsZ gene (Kelly et al., 1998), repetitive extragenic palindromic (REP) or enterobacterial repetitive intergenic consensus (ERIC) DNA sequences (Clarridge et al., 1995; Rodriguez-Barradas et al., 1995b) and the 16S-23S rDNA intergenic spacer (ITS; Minnick and Barbian, 1997b).

Restriction Fragment-Length Polymorphism

Fig. 2. Transmission electron micrographs showing the two basic cell morphologies observed in Bartonella species. Bacteria were grown by standard methods and stained with uranyl acetate. A) Flagellated cell morphology as exemplified by B. clarridgeiae, and B) a more typical, nonflagellated cell morphology as exemplified by B. elizabethae (Bars = 0.5 µm). (Courtesy of James A. Carroll.)

The RFLPs identifying Bartonella spp. employed rare-cutting restriction endonucleases together with genomic DNA (Slater et al., 1990; Maurin et al., 1994; Roux and Raoult, 1995), with PCR fragments of gltA (Norman et al., 1995), or with PCR products containing the 16S-23S rDNA intergenic spacer region (ITS; Matar et al., 1993; Roux and Raoult, 1995; Bergmans et al., 1996).

Cellular Fatty Acids Cellular fatty acid (CFA) analysis also has been used to identify Bartonella spp. to the genus level. This technique is particularly useful, as Bar-

CHAPTER 3.1.18

tonella spp. have an unusual fatty acid composition when compared to other bacteria. cis-11-Octadecanoate (C18:1 ω7c; ~54%) and hexadecanoate (C16:0; ~20%) are the predominant fatty acids of all Bartonella spp. Bartonella elizabethae, B. henselae, B. clarridgeiae and B. quintana contain considerable amounts of octadecanoate (C18:0; ~23%), whereas B. bacilliformis contains very little (~2%). Bartonella bacilliformis also contains an unusually high quantity (~20%) of cis-11 hexadecanoate (C16:1 ω7c) in contrast to most Bartonella species (64 by immunofluorescence assay is considered positive for B. henselae by the United States Centers for Disease Control and Prevention (CDC; Regnery et al., 1992b). When available, PCR on lymph node aspirates or biopsies is often successful (Anderson et al., 1994b). BACILLARY ANGIOMATOSIS. Patients usually are immunocompromised. Cutaneous and/or subcutaneous nongranulomatous vascular lesions affect a variety of organs. Bacilli in sections of affected tissue can be visualized using Warthin-Starry silver stain. Diagnostic testing includes an anti-B. henselae IgG indirect fluorescent antibody assay (Regnery et al., 1992a; Zangwill et al., 1993; Patnaik et al., 1992) and an ELISA system (Barka et al., 1993). BACILLARY PELIOSIS. Patients usually are immunocompromised. Diagnosis based on

The Genus Bartonella

483

symptoms is difficult and may require biopsy. Cystic blood-filled liver lesions that are similar to those of BA, together with bacilli in sections of affected tissue, are diagnostic. Hepatomegaly and elevated serum levels of liver enzymes are characteristic. Bacillary peliosis should be suspected if patient has BA or bacteremic syndrome. Detection reagents used for BA diagnosis also can be used for bacillary peliosis. OROYA FEVER. Patients have a history of travel or residence in South America and may have history of sandfly bite. Diagnostic symptoms of the primary stage include a dramatically low erythrocyte count (~500,000/mm3; Hurtado et al., 1938; Reynafarje and Ramos, 1961; Kreier and Ristic, 1981) and numerous infected erythrocytes when blood smears are stained with eosin and thiazine (Knobloch et al., 1985). Secondary (tissue) phase presents with cutaneous angiomatous lesions (verruga peruana) that contain bacteria when sectioned and stained with WarthinStarry silver stain. TRENCH FEVER. Patients likely have a history of infestation with body lice (Pediculus humanus). Chronic alcohol abuse, homelessness and non-Caucasian descent are risk factors (Jackson and Spach, 1996). Diagnostic characteristics include pain in the bones (especially the tibia), fever and persistent bacteremia for several weeks to months following resolution of symptoms (Vinson et al., 1969). The presence of seroconversion to B. quintana can be tested; however, the antibody response usually is not specific to only B. quintana owing to crossreactivity within the genus (Regnery et al., 1992a). ENDOCARDITIS. Bartonella infection should be considered in blood culture-negative cases of endocarditis. Patients usually have predisposing heart conditions or are at high risk for contracting infection with B. henselae or B. quintana (see B. henselae or B. quintana under “Epidemiology and Control”). Vegetative lesions are common in cases of Bartonella endocarditis (Daly et al., 1993; Raoult et al., 1996). The difficulty of culture from patient isolates and serological crossreactivity between Chlamydia spp., Bartonella spp. and Coxiella burnetii can complicate diagnosis (Drancourt et al., 1995; Knobloch et al., 1988b; La-Scola and Raoult, 1996). Seropositivity for Bartonella infection can be tested with the detection reagents described for diagnosis of BA. Molecular techniques involving PCR also should be used, particularly if species-level identification is desired (see “Identification”). BACTEREMIC SYNDROME. Patients are usually immunocompromised. Persistent bacteremia and recurring fevers, together with chronic fatigue and malaise, are diagnostic. Diagnosis can be made by blood cultures with subsequent identification protocols (see “Identification”) or

484

M.F. Minnick and B.E. Anderson

CHAPTER 3.1.18

by testing for seropositivity to B. henselae or B. quintana as described for BA.

Antimicrobial Therapy A variety of antibiotics have been used successfully to treat bartonelloses. Antimicrobial therapy varies depending upon Bartonella species and in some cases is syndrome-dependent. CSD and trench fever generally are self-limiting illnesses that do not require therapy unless the infection is systemic. In contrast, BA can be life threatening and requires prompt antibiotic therapy. The immune state of the patient is also an important consideration for prescribing the type and duration of antimicrobial therapy for bartonellosis. For example, BA therapy for healthy patients is approximately three weeks, whereas immunocompromised patients may require several weeks to months, or possibly a lifetime, of therapy to resolve the infection (Adal et al., 1994). Relapse of bartonellosis is common even with antimicrobial therapy. Bartonellae are typically sensitive to most antibiotics in vitro (Maurin and Raoult, 1996), but are frequently resistant to them during human therapy. This observation may reflect the inability of the chemotherapeutic agent to access the pathogen located within host cells. Aminoglycosides are particularly bactericidal for bartonellae in vivo (Musso et al., 1995) and are recommended for therapy. Although optimal antibiotic regimens are still unclear, a list of antimicrobials that have successfully been used in treating various manifestations of bartonellosis is given in Table 4.

Immunity The general course of bartonellosis begins with transient or persistent bacteremia that may involve erythrocyte parasitism, depending upon bacterial species. The pathogen subsequently infects the vascular bed within a variety of tissues, where vascular lesions are produced. Tissue involvement can be chronic, and presumably the pathogen is shed from the affected tissues into the bloodstream. Lymphadenopathy is a frequent complication in bartonellosis. The immune status of the host is a risk factor for infection. Typically, immunocompromised patients are at higher risk of opportunistic infection by bartonellosis than healthy individuals are. As a rule of thumb, trench fever, CSD, endocarditis and Oroya fever take place regardless of immune status of the patient, whereas BA, bacillary peliosis and bacteremic syndrome occur mainly in immuocompromised individuals. Bartonellosis symptoms are generally more pronounced in immunodeficient patients. In cases of BA, many patients display a marked CD4+ lymphocyte leukocytopenia, with counts of less than 100 (LeBoit et al., 1989; Koehler and Tappero, 1993). At best, the human immune response against Bartonella is poorly characterized, and for some species (B. elizabethae and B. clarridgeiae) it has never been studied. Because Bartonella spp. are found in a variety of body fluids and cells, involvement of both humoral and cellular arms of the immune system needs to be invoked for resolution of infection. BARTONELLA BACILLIFORMIS. Lifelong humoral immunity supposedly results from an

Table 4. Antimicrobial agents used to treat bartonelloses. Syndrome BA

Bacteremic syndrome BP CSDb

Endocarditis Oroya fever Pulmonary nodules Retinal vasculitis and vitreitis Trench fevera

Antimicrobial Agent(s) Erythromycin or doxycycline Also chloramphenicol, sulfa-trimethoprim, aminoglycosides, azithromycin Ciprofloxacin Clarithromicin Erythromycin or doxycycline Erythromycina Ciprofloxacin, sulfa-trimethoprim or gentamicin Erythromycin or doxycycline Azithromycin Nafcillin + gentamicin β-Lactams, tetracycline + streptomycin, aminoglycosides, chloramphenicol if coninfected by enterics Doxycycline Ciprofloxacin Tetracycline or chloramphenicol

Abbreviations: BA, bacillary angiomatosis; BP, bacillary peliosis; and CSD, cat-scratch disease. Alternatives as in BA. b Usually self-limiting (does not require antibiotic therapy unless systemic). a

Reference(s) Bartlett, 1996 Milde et al., 1995

Adal et al., 1994 Perkocha et al., 1990 Bartlett, 1996 Bartlett, 1996 Smith, 1997 Bass, 1998 Daly et al., 1993 Weinman, 1965 Caniza et al., 1995 Soheilian et al., 1996 Moulder, 1974

CHAPTER 3.1.18

active infection with B. bacilliformis (Ricketts, 1949; Weinman, 1965). Seropositivity (mainly IgM isotype) in endemic areas of Peru can reach 60% of the population, and many of these individuals are healthy (Knobloch et al., 1985). However, in spite of circulating antibody, many individuals who are asymptomatic or posteruptive for verruga peruana are often blood culturepositive for B. bacilliformis (Howe, 1943), and carrier rates can reach 15% in some areas of Peru (Herrer, 1953). Immunosuppression is characteristic and probably results from the acute anemia and leukocytosis. Although a variety of proteins from B. bacilliformis are immunogenic (Knobloch, 1988a), the most prominent antigens include a GroEL homologue (BB65) (Knobloch and Schreiber, 1990), the flagellin protein (42 kDa), a phage coat protein (31.5 kDa) and an uncharacterized protein of 45 kDa (Minnick, 1994). The gene encoding a 43-kDa lipoprotein of B. bacilliformis reactive with human sera from bartonellosis cases was recently cloned, sequenced and expressed (Padmalayam et al., 2000b). BARTONELLA QUINTANA. Although symptoms during B. quintana infections are highly variable, patients generally produce lowtiter, complement-fixing antibody during the early phase of convalescence (Varela et al., 1969). However, in spite of antibodies, patients frequently display persistent and protracted bacteremia (Varela et al., 1969; Koehler et al., 1992; Maurin et al., 1994). In addition, B. quintana is markedly resistant to killing by complement fixed by classical or alternative pathways (Myers and Wisseman, 1978). Infection frequently is accompanied by leukocytosis (Vinson et al., 1969), possibly enhancing bacteremia. BARTONELLA HENSELAE. Patients infected with B. henselae are generally seropositive when assayed by indirect fluorescence antibody tests (Regnery et al., 1992a). Antibodies apparently opsonize the pathogen and enhance production of oxygen radicals following phagocytosis, but do not enhance complement activation by the classical pathway. In fact, complement fixation mainly proceeds via the alternative pathway (Rodriguez-Barradas et al., 1995a). Persistent bacteremia with B. henselae is common in both humans and cats (Koehler et al., 1994), and respective antibody titers do not necessarily correlate with protection against the pathogen. A 17-kDa antigen has been shown to be highly reactive with human sera from patients with CSD (Anderson et al., 1995). The gene encoding this antigen is conserved among most Bartonella spp. but shows extensive amino acid sequence divergence (Sweger et al., 2000). A second protein of 83-kDa has been reported to react with human sera from patients with CSD

The Genus Bartonella

485

(McGill et al., 1998). In addition to antibody reactivity, the cellular immune response in humans with CSD has been well recognized (Gerber et al., 1986). The aggravating cellular immune response to B. henselae both contributes to the pathogenesis of CSD and forms the basis of diagnostic skin test. This delayed-type hypersensitivity and T helper 1-type cytokine response has been recently reproduced in mouse models (Arvand et al., 1998; Karem et al., 1999).

Literature Cited Adal, K. A., C. J. Cockerell, and W. A. Petri. 1994. Cat scratch disease, bacillary angiomatosis and other infections due to Rochalimaea. N. Engl. J. Med. 330:1509–1515. Alexander, B. 1995. A review of bartonellosis in Ecuador and Colombia. Am. J. Trop. Med. Hyg. 52:354–359. Anderson, B., C. Kelley, R. Threlkel, and K. Edwards. 1993. Detection of Rochalimaea henselae in catscratch disease skin test antigens. J. Infect. Dis. 168:1034–1036. Anderson, B., C. Goldsmith, A. Johnson, I. Padmalayam, and B. Baumstark. 1994a. Bacteriophage-like particle of Rochalimaea henselae. Molec. Microbiol. 13:67–73. Anderson, B., K. Sims, R. Regnery, L. Robinson, M. J. Schmidt, S. Gorel, C. Hager, and K. Edwards. 1994b. Detection of Rochalimaea henselae DNA in clinical specimens from cat-scratch disease patients by using polymerase chain reaction. J. Clin. Microbiol. 32:942– 948. Anderson, B., E. Lu, D. Jones, and R. Regnery. 1995. Characterization of a 17-kDa antigen of Bartonella henselae reactive with sera from patients with cat-scratch disease. J. Clin. Microbiol. 33:2358–2365. Anderson, B., D. Jones, and A. Burgess. 1996. Cloning, expression and sequence analysis of the Bartonella henselae gene encoding the HtrA sress-response protein. Gene 178:35–38. Anderson, B., D. Scotchlas, D. Jones, A. Johnson, T. Tzianabos, and B. Baumstark. 1997b. Analysis of 36-kilodalton protein (PapA) associated with the bacteriophage particle of Bartonella henselae. DNA Cell Biol. 16:1223– 1229. Angritt, P., S. M. Tuur, A. M. Macher, K. J. Smith, C. S. Park, F. P. Hobin, and C. Myrie-Williams. 1988. Epithelioid angiomatosis in HIV infection: Neoplasm or cat-scratch disease? Lancet 1:996. Arias-Stella, J., P. H. Lieberman, R. A. Erlandson, and J. Arias-Stella Jr. 1986. Histology, immunohistochemistry, and ultrastructure of the verruga in Carrion’s disease. Am. J. Surg. Pathol. 10:595–610. Barbian, K. D., and M. F. Minnick. 2000. A bacteriophagelike particle from Bartonella bacilliformis. Microbiology 146:599–609. Barka, N. E., T. Hadfield, M. Patnaik, W. A. Schwartzman, and J. B. Peter. 1993. EIA for detection of Rochalimaea henselae-reactive IgG, IgM and IgA antibodies in patients with suspected cat-scratch disease. J. Infect. Dis. 167:1503–1504. Bartlett, J. G. 1996. Pocket Book of Infectious Disease Therapy. Williams and Wilkins Baltimore, MD. 22. Barton, A. 1909. Descripcion de elementos endoglobulares en los enfermos de Fiebre de Verruga. Cron. Med. Lima 26:7.

486

M.F. Minnick and B.E. Anderson

Bass, J. W., B. C. Freitas, A. D. Freitas, C. L. Sisler, D. S. Chan, J. M. Vincent, D. A. Person, J. R. Claybaugh, R. R. Wittler, M. E. Weisse, R. L. Regnery, and L. N. Slater. 1998. Prospective randomized double blind placebocontrolled evaluation of azithromycin for treatment of cat-scratch disease. Pediatr. Infect. Dis. J. 17:447–452. Batterman, H. J., J. A. Peek, J. S. Loutit, S. Falkow, and L. S. Tompkins. 1995. Bartonella henselae and Bartonella quintana adherence to and entry into cultured human epithelial cells. Infect. Immun. 63:4553–4556. Battisti, J. M., L. S. Smitherman, D. S. Samuels, and M. F. Minnick. 1998. Mutations in Bartonella bacilliformis gyrB confer resistance to coumermycin A1. Antimicrob. Agents Chemother. 42:2906–2913. Battisti, J. M., and M. F. Minnick. 1999. Development of a system for genetic manipulation of Bartonella bacilliformis. Appl. Environ. Microbiol. 65:3441–3448. Baumler, A. J., J. G. Kusters, I. Stojiljkovic, and F. Heffron. 1994. Salmonella typhimurium loci involved in survival within macrophages. Infect. Immun. 62:1623–1630. Benson, L. A., S. Kar, G. McLaughlin, and G. M. Ihler. 1986. Entry of Bartonella bacilliformis into erythrocytes. Infect. Immun. 54:347–353. Bereswill, S., S. Hinklemann, M. Kist, and A. Sander. 1999. Molecular analysis of riboflavin synthesis genes in Bartonella henselae and use of the ribC gene for differentiation of Bartonella species by PCR. J. Clin. Microbiol. 37:3159–3166. Berger, T. G., J. W. Tappero, A. Kaymen, and P. E. LeBoit. 1989. Bacillary (epithelioid) angiomatosis and concurrent Kaposi’s sarcoma in acquired immunodeficiency syndrome. Arch. Dermatol. 125:1543–1547. Bergmans, A. M., J. W. Groothedde, J. F. Schellekens, J. D. van Embden, J. M. Ossewaarde, and L. M. Schouls. 1995. Etiology of cat scratch disease: Comparison of polymerase chain reaction detection of Bartonella (formerly Rochalimaea) and Afipia felis DNA with serology and skin tests. J. Infect. Dis. 171:916–23. Bergmans, A. M., J. F. Schellekens, J. D. VanEmbden, and L. M. Schouls. 1996. Predominance of two Bartonella henselae variants among cat-scratch disease patients in the Netherlands. J. Clin. Microbiol. 34:254–260. Birtles, R. J., T. G. Harrison, N. K. Fry, N. A. Saunders, and A. G. Taylor. 1991. Taxonomic considerations of Bartonella bacilliformis based on phylogenetic and phenotypic characteristics. FEMS Microbiol. Lett. 67:187–191. Birtles, R. J., T. G. Harrison, N. A. Saunders, and D. H. Molyneux. 1995. Proposals to unify the genera Grahamella and Bartonella, with descriptions of Bartonella talpae comb. nov., Bartonella peromysci comb. nov., and three new species, Bartonella grahamii sp. nov., Bartonella taylorii sp. nov., and Bartonella doshiae sp. nov. Int. J. Syst. Bacteriol. 45:1–8. Birtles, R. J., and D. Raoult. 1996. Comparison of partial citrate synthase gene (gltA) sequences for phylogenetic analysis of Bartonella species. Int. J. Syst. Bacteriol. 46:891–897. Bowers, T. J., D. Sweger, D. Jue, and B. Anderson. 1998. Isolation, sequencing and expression of the gene encoding a major protein from the bacteriophage associated with Bartonella henselae. Gene 206:49–52. Breitschwerdt, E. B., and D. L. Kordick. 2000. Bartonella infection in animals: Carriership, reservoir potential, pathogenicity, and zoonotic potential for human infection. Clin. Microbiol. Rev. 13(3):428–438.

CHAPTER 3.1.18 Brenner, D. J., S. P. O’Connor, D. G. Hollis, R. E. Weaver, and A. G. Steigerwalt. 1991. Molecular characterization and proposal of a neotype strain for Bartonella bacilliformis. J. Clin. Microbiol. 29:1299–1302. Brenner, D. J., S. P. O’Connor, H. H. Winkler, and A. G. Steigerwalt. 1993. Proposals to unify the genera Bartonella and Rochalimaea, with descriptions of Bartonella quintana comb. nov., Bartonella vinsonii comb. nov., Bartonella henselae comb. nov., and Bartonella elizabethae comb. nov., and to remove the family Bartonellaceae from the order Rickettsiales. Int. J. Syst. Bacteriol. 43:777–786. Brouqui, P., and D. Raoult. 1996. Bartonella quintana invades and multiplies within endothelial cells in vitro and in vivo and forms intracellular blebs. Res. Microbiol. 147:719–731. Burgess, W. O., and B. E. Anderson. 1998. Outer membrane proteins of Bartonella henselae and their interaction with human endothelial cells. Microb. Pathog. 25:157– 164. Byam, W. 1919. Trench fever. In: L. L. Loyd (Ed.) Lice and Their Menace to Man. Oxford University Press. Oxford, 120–130. Caniza, M. A., D. L. Granger, K. H. Wilson, M. K. Washington, D. L. Kordick, D. P. Frush, and R. B. Blitchington. 1995. Bartonella (Rochalimaea) henselae: Etiology of pulmonary nodules in a patient with depressed cellmediated immunity. Clin. Infect. Dis. 20:1505–1511. Carithers, H. A. 1985. Cat-scratch disease: An overview based on a study of 1,200 patients. Am. J. Dis. Child. 139:1124–1133. Carroll, J. A., S. A. Coleman, L. S. Smitherman, and M. F. Minnick. 2000. Hemin-binding surface protein from Bartonella quintana. Infect. Immun. 68:6750–6757. Cartwright, J. L., P. Britton, M. F. Minnick, and A. G. McLennan. 1999. The ialA invasion gene of Bartonella bacilliformis encodes a (di)nucleoside polyphosphate hydrolase of the MutT motif family and has homologs in other invasive bacteria. Biochem. Biophys. Res. Comm. 256:474–479. Chomel, B. B., R. W. Kasten, K. Floyd-Hawkins, B. Chi, K. Yamamoto, J. Roberts-Wilson, A. N. Gurfield, R. C. Abbott, N. C. Pedersen, and J. E. Koehler. 1996. Experimental transmission of Bartonella henselae by the cat flea. J. Clin. Microbiol. 34:1952–1956. Clarridge, J. E., T. J. Raich, D. Pirwani, B. Simon, L. Tsai, M. C. Rodriguez-Barradas, R. Regnery, A. Zollo, D. C. Jones, and C. Rambo. 1995. Strategy to detect and identify Bartonella species in routine clinical laboratory yields Bartonella henselae from human immunodeficiency virus-positive patient and unique Bartonella strain from his cat. J. Clin. Microbiol. 33:2107–2113. Cockerell, C. J., M. A. Whitlow, G. F. Webster, and A. E. Friedman-Kien. 1987. Epithelioid angiomatosis: A distinct vascular disorder in patients with the acquired immunodeficiency syndrome or AIDS-related complex. Lancet 2:654–656. Cockerell, C. J., P. R. Bergstresser, C. Myrie-Williams, and P. M. Tierno. 1990. Bacillary epithelioid angiomatosis occurring in an immunocompetent individual. Arch. Dermatol. 126:787–790. Conley, T., L. Slater, and K. Hamilton. 1994. Rochalimaea species stimulate human endothelial cell proliferation and migration in vitro. J. Lab. Clin. Med. 124:521–528. Cuadra, M. S. 1956. Salmonellosis complication in human bartonellosis. Tex. Rep. Biol. Med. 14:97–113.

CHAPTER 3.1.18 Daly, J. S., M. G. Worthington, D. J. Brenner, C. W. Moss, D. G. Hollis, R. S. Weyant, A. G. Steigerwalt, R. E. Weaver, M. I. Daneshvar, and S. P. O’Connor. 1993. Rochalimaea elizabethae sp. nov. isolated from a patient with endocarditis. J. Clin. Microbiol. 31:872–881. Dauga, C., I. Miras, P. A. Grimont. 1996. Identification of Bartonella henselae and B. quintana 16S rDNA sequences by branch-, genus- and species-specific amplification. J. Med. Microbiol. 45:192–199. Dehio, C., and M. Meyer. 1997a. Maintenance of broadhost-range incompatibility group P and group Q plasmids and transposition of Tn5 in Bartonella henselae following conjugal transfer from Escherichia coli. J. Bacteriol. 179:538–540. Dehio, C., M. Meyer, J. Berger, H. Schwarz, and C. Lanz. 1997b. Interaction of Bartonella henselae with endothelial cells results in bacterial aggregation on the cell surface and the subsequent engulfment and internalisation of the bacterial aggregate by a unique structure, the invasome. J. Cell Sci. 110:2141–2154. Dolan, M. J., M. T. Wong, R. L. Regnery, J. H. Jorgensen, M. Garcia, J. Peters, and D. Drehner. 1993. Syndrome of Rochalimaea henselae adenitis suggesting cat scratch disease. Ann. Intern. Med. 118:331–336. Doyle, D., S. C. Eppes, and J. D. Klein. 1994. Atypical catscratch disease: Diagnosis by a serologic test for Rochalimaea species. South. Med. J. 87:485–487. Drancourt, M., and D. Raoult. 1993. Proposed tests for the routine identification of Rochalimaea species. Eur. J. Clin. Microbiol. Infect. Dis. 12:710–713. Drancourt, M., J. L. Mainardi, P. Brouqui, F. Vandenesch, A. Carta, F. Lehnert, J. Etienne, F. Goldstein, J. Acar, and D. Raoult. 1995. Bartonella (Rochalimaea) quintana endocarditis in three homeless men. N. Engl. J. Med. 332:419–423. Drancourt, M., R. Birtles, G. Chaumentin, F. Vandenesch, J. Etienne, and D. Raoult. 1996. New serotype of Bartonella henselae in endocarditis and cat-scratch disease. Lancet 347:441–443. Droz S., B. Chi, E. Horn, A. G. Steigerwalt, A. M. Whitney, and D. J. Brenner. 1999. Bartonella koehlerae sp. nov., isolated from cats. J. Clin. Microbiol. 37:1117–1122. Edmond, M. B., S. A. Riddler, C. M. Baxter, B. M. Wicklund, and A. W. Pasculle. 1993. Agrobacterium radiobacter: A recently recognized opportunistic pathogen. Clin. Infect. Dis. 16:388–391. Ehrenborg, C., L. Wesslen, A. Jakobson, G. Friman, and M. Holmberg. 2000. Sequence variation in the ftsZ gene of Bartonella henselae isolates and clinical samples. J. Clin. Microbiol. 38:682–687. English, C. K., D. J. Wear, A. M. Margileth, C. R. Lissner, and G. P. Walsh. 1988. Cat-scratch disease. Isolation and culture of the bacterial agent. JAMA 259:1347–1352. Fath, M. J., and R. Kolter. 1993. ABC transporters: Bacterial exporters. Microbiol. Rev. 57:995–1017. Fischetti, V. A. 1989. Streptococcal M protein: Molecular design and biological behavior. Clin. Microbiol. Rev. 2:285–314. Freney, J., L. D. Gruer, N. Bornstein, M. Kiredjian, I. Guilvout, M. N. Letouzey, C. Combe, and J. Fleurette. 1985. Septicemia caused by Agrobacterium sp. J. Clin. Microbiol. 22:683–685. Garcia, F. U., J. Wojta, K. N. Broadley, J. M. Davidson, and R. L. Hoover. 1990. Bartonella bacilliformis stimulates endothelial cells in vitro and is angiogenic in vivo. Am. J. Pathol. 136:1125–1135.

The Genus Bartonella

487

Garcia, F. U., J. Wojta, and R. L. Hoover. 1992. Interactions between live Bartonella bacilliformis and endothelial cells. J. Infect. Dis. 165:1138–1141. Garcia-Tsao, G., L. Panzini, M. Yoselevitz, and A. B. West. 1992. Bacillary peliosis hepatis as a cause of acute anemia in a patent with the acquired immunodeficiency syndrome. Gastroenterol. 102:1065–1070. Gerber, M. A., P. Rapacz, S. S. Kalter, and M. Ballow. 1986. Cell-mediated immunity in catscratch disease. J. Allergy Clin. Immunol. 78:887–890. Gille, C., C. Lanz, and C. Dehio. 1999. Site-specific mutagenesis of the ialB locus of Bartonella tribocorum by singlecrossover gene disruption. In: Abstracts of the First International Conference on Bartonella as Emerging Pathogens. Max Planck Institute. Tuebingen, Germany. 56, poster 28. Gold, L., D. Pribnow, T. Schneider, S. Shinedling, B. S. Singer, and G. Stormo. 1981. Translation initiation in prokaryotes. Ann. Rev. Microbiol. 35:365–403. Golnik, K. C., M. E. Marotto, M. M. Fanous, D. Heitter, L. P. King, J. I. Halpern, and P. H. Holly. 1994. Opthalmic manifestations of Rochalimaea species. Am. J. Opthalmol. 118:145–151. Gray, G. C., A. A. Johnson, S. A. Thornton, W. A. Smith, J. Knobloch, P. W. Kelley, L. O. Escudero, M. A. Huayda, and F. S. Wignall. 1990. An epidemic of Oroya fever in the Peruvian Andes. Am. J. Trop. Med. Hyg. 42:215–221. Gurfield, A. N., H. J. Boulouis, B. B. Chomel, R. Heller, R. W. Kasten, K. Yamamoto, and Y. Piemont. 1997. Coinfection with Bartonella clarridgeiae and Bartonella henselae and with different Bartonella henselae strains in domestic cats. J. Clin. Microbiol. 35:2120–2123. Haake, D. A., T. A. Summers, A. M. McCoy, and W. Schwartzman. 1997. Heat shock response and groEL sequence of Bartonella henselae and Bartonella quintana. Microbiol. 143:2807–2815. Hadfield, T. L., R. Warren, M. Kass, E. Brun, and L. Levy. 1993. Endocarditis caused by Rochalimaea henselae. Hum. Pathol. 24:1140–1141. Heffernan, E. J., L. Wu, J. Louie, S. Okamoto, J. Fierer, and D. G. Guiney. 1994. Specificity of the complement resistance and cell association phenotypes encoded by the outer membrane protein genes rck from Salmonella typhimurium and ail from Yersinia enterocolitica. Infect. Immun. 62:5183–5186. Heller, R., P. Riegel, Y. Hansmann, G. Delacour, D. Bermond, C. Dehio, F. Lamarque, H. Monteil, B. Chomel, and Y. Piemont. 1998. Bartonella tribocorum sp. nov. a new Bartonella species isolated from the blood of wild rats. Int. J. Syst. Bacteriol. 48:1333–1339. Heller, R., M. Kubina, P. Mariet, P. Riegel, G. Delacour, C. Dehio, F. Lamarque, R. Kasten, H. J. Boulouis, H. Monteil, B. Chomel, and Y. Piemont. 1999. Bartonella alsatica sp. nov., a new Bartonella species isolated from the blood of wild rabbits. Int. J. Syst. Bacteriol. 49:283–288. Herrer, A. 1953. Carrion’s disease. II: Presence of Bartonella bacilliformis in the peripheral blood of patients with the benign tumor form. Am. J. Trop. Med. 2:645–649. Hertig, M. 1942. Phlebotomus and Carrion’s disease. Am. J. Trop. Med. 22:1–80. Hollingdale, M. R., J. W. Vinson, and J. E. Herrmann. 1980. Immunochemical and biological properties of the outer membrane-associated lipopolysaccharide and protein of Rochalimaea quintana. J. Infect. Dis. 141:672–679. Holmes, A., T. Greenough, G. Balady, R. Regnery, B. Anderson, J. O’Keane, and E. McCrone. 1995. Bartonella

488

M.F. Minnick and B.E. Anderson

henselae endocarditis in an immunocompetent adult. Clin. Infect. Dis. 21:1004–1007. Howe, C. 1943. Carrion’s disease: Immunologic studies. Arch. Intern. Med. 72:147–167. Hurtado, A., J. P. Musso, and C. Merino. 1938. La anemia en la enfermedad de Carrion (verruga peruana). Ann. Fac. Med. Lima 28:154–168. Iwaki-Egawa S., and G. M. Ihler. 1997. Comparison of the abilities of proteins from Bartonella bacilliformis and Bartonella henselae to deform red cell membranes and to bind to red cell ghost proteins. FEMS Microbiol. Lett. 157:207–217. Jackson, L. A., and D. H. Spach. 1996. Emergence of Bartonella quintana infection among homeless persons. Emerg. Infect. Dis. 2:141–144. Jacobs, R. F., and G. E. Schutze. 1998. Bartonella henselae as a cause of prolonged fever and fever of unknown origin in children. Clin. Infect. Dis. 26:80–84. Jalava J., P. Kotilainen, S. Nikkari, M. Skurnik, E. Vanttinen, O. P. Lehtonen, E. Eerola, and P. Toivanen. 1995. Use of the polymerase chain reaction and DNA sequencing for detection of Bartonella quintana in the aortic valve of a patient with culture-negative infective endocarditis. Clin. Infect. Dis. 21:891–896. Johnson, W. T., and E. B. Helwig. 1969. Cat-scratch disease: Histopathologic changes in the skin. Arch. Dermatol. 100:148–154. Karem, K. L., K. A. Dubois, S. L. McGill, and R. L. Regnery. 1999. Characterization of Bartonella henselae-specific immunity in BALB/c mice. Immunology 97:352–358. Kehoe, M. A. 1994. Cell wall-associated proteins in Grampositive bacteria. New Compr. Biochem. 27:217–261. Kelly, T. M., I. Padmalayam, and B. R. Baumstark. 1998. Use of the cell division protein FtsZ as a means of differentiating among Bartonella species. Clin. Diagn. Lab. Immunol. 5:766–772. Kemper, C. A., C. M. Lombard, S. C. Deresinski, and L. S. Tompkins. 1990. Visceral bacillary epithelioid angiomatosis: Possible manifestations of disseminated cat scratch disease in the immunocompromised host. A report of two cases. Am. J. Med. 89:216–222. Kerkhoff, F. T., A. M. Bergmans, A. van Der Zee, and A. Rothova. 1999. Demonstration of Bartonella grahamii DNA in ocular fluids of a patient with neuroretinitis. J. Clin. Microbiol. 37:4034–4038. Knobloch, J., L. Solano, O. Alvarez, and E. Delgado. 1985. Antibodies to Bartonella bacilliformis as determined by fluorescence antibody test, indirect haemagglutination and ELISA. Trop. Med. Parasitol. 36:183–185. Knobloch, J. 1988a. Analysis and preparation of Bartonella bacilliformis antigens. Am. J. Trop. Med. Hyg. 39:173– 178. Knobloch, J., R. Bialek, G. Muller, and P. Asmus. 1988b. Common surface epitope of Bartonella bacilliformis and Chlamydia psittaci. Am. J. Trop. Med. Hyg. 39:427–433. Knobloch, J., and M. Schreiber. 1990. BB65, a major immunoreactive protein of Bartonella bacilliformis. Am. J. Trop. Med. Hyg. 43:373–379. Kobe, B., and J. Deisenhofer. 1995. Proteins with leucine-rich repeats. Curr. Opin. Struct. Biol. 5:409–416. Koehler, J. E., F. D. Quinn, T. G. Berger, P. E. LeBoit, and J. W. Tappero. 1992. Isolation of Rochalimaea species from cutaneous and osseous lesions of bacillary angiomatosis. N. Engl. J. Med. 327:1625–1631. Koehler, J. E., and J. W. Tappero. 1993. Bacillary angiomatosis and bacillary peliosis in patients infected with

CHAPTER 3.1.18 human immunodeficiency virus. Clin. Infect. Dis. 17:612–624. Koehler, J. E., C. A. Glaser, and J. W. Tappero. 1994. Rochalimaea henselae infection: A new zoonosis with the domestic cat as reservoir. JAMA 271:531–535. Kordick, D. L., and E. B. Breitschwerdt. 1995a. Intaerythrocytic presence of Bartonella henselae. J. Clin. Microbiol. 33:1655–1656. Kordick, D. L., K. H. Wilson, D. J. Sexton, T. L. Hadfield, H. A. Berkhoff, and E. B. Breitschwerdt. 1995b. Prolonged Bartonella bacteremia in cats associated with catscratch disease patients. J. Clin. Microbiol. 33:3245–3251. Kordick, D. L., B. Swaminathan, C. E. Greene, K. H. Wilson, A. M. Whitney, S. O’Connor, D. G. Hollis, G. M. Matar, A. G. Steigerwalt, G. B. Malcolm, P. S. Hayes, T. L. Hadfield, E. B. Breitschwerdt, and D. J. Brenner. 1996. Bartonella vinsonii subsp. berkhoffii subsp.nov., isolated from dogs; Bartonella vinsonii subsp. vinsonii; and emended description of Bartonella vinsonii. Int. J. Syst. Bacteriol. 46:704–709. Kordick, D. L., E. J. Hilyard, T. L. Hadfield, K. H. Wilson, A. G. Steigerwalt, D. J. Brenner, and E. B. Breitschwerdt. 1997. Bartonella clarridgeiae, a newly recognized zoonotic pathogen causing inoculation papules, fever, and lymphadenopathy (cat scratch disease). J. Clin. Microbiol. 35:1813–1818. Kostrzewski, J. 1950. The epidemiology of trench fever. Med. Dosw. Mikrobiol. 11:233–263. Kreier, J. P., and M. Ristic. 1981. The biology of hemotrophic bacteria. Ann. Rev. Microbiol. 35:325–338. Kreier, J. P., R. Gother, G. M. Ihler, H. E. Krampitz, G. Mernaugh, and G. H. Palmer. 1991. The hemotrophic bacteria: The families Bartonellaceae and Anaplasmataceae. In: A. Balows, H. G. Truper, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York, NY. 3994– 4022. Krueger, C. M., K. L. Marks, and G. M. Ihler. 1995. Physical map of the Bartonella bacilliformis genome. J. Bacteriol. 177:7271–7274. La-Scola, B., and D. Raoult. 1996. Serological cross-reactions between Bartonella quintana, Bartonella henselae, and Coxiella burnetii. J. Clin. Microbiol. 34:2270–2274. LeBoit, P. E., T. G. Berger, B. M. Egbert, J. H. Beckstead, T. S. Yen, and M. H. Stoler. 1988. Epithelioid haemangioma-like vascular proliferation in AIDS: Manifestation of cat-scratch disease bacillus infection? Lancet 1:960–963. LeBoit, P. E., T. G. Berger, B. M. Egbert, J. H. Beckstead, T. S. Yen, and M. H. Stoler. 1989. Bacillary angiomatosis. The histopathology and differential diagnosis of a pseudoneoplastic infection in patients with human immunodeficiency virus disease. Am J. Surg. Pathol. 13:909–920. Lee, A. K., and S. Falkow. 1998. Constitutive and inducible green fluorescent protein expression in Bartonella henselae. Infect. Immun. 66:3964–3967. Lucey, D., M. J. Dolan, C. W. Moss, M. Garcia, D. G. Hollis, S. Wegner, G. Morgan, R. Almeida, D. Leong, K. S. Greisen, D. F. Welch, and L. N. Slater. 1992. Relapsing illness due to Rochalimaea henselae in immunocompetent hosts: Implication for therapy and new epidemiological associations. Clin. Infect. Dis. 14:683– 688. Maeno, N., H. Oda, K. Yoshiie, M. R. Wahid, T. Fujimura, and S. Matayoshi. 1999. Live Bartonella henselae

CHAPTER 3.1.18 enhances endothelial cell proliferation without direct contact. Microb. Path. 27:419–427. Margileth, A. M. 1993. Cat scratch disease. Adv. Pediatr. Infect. Dis. 8:1–21. Margileth, A. M., and D. F. Baehren. 1998. Chest-wall abscess due to cat-scratch disease (CSD) in an adult with antibodies to Bartonella clarridgeiae: Case report and review of the thoracopulmonary manifestations of CSD. Clin. Infect. Dis. 27:353–357. Marullo, S., A. Jaccard, D. Roulot, C. Mainquene, and J. P. Clauvel. 1992. Identification of the Rochalimaea henselae 16S rRNA sequence in the liver of a French patient with bacillary peliosis hepatis. J. Infect. Dis. 166:1462. Maruyama, S., S. Nogami, I. Inoue, S. Namba, K. Asanome, and Y. Katsube. 1996. Isolation of Bartonella henselae from domestic cats in Japan. J. Vet. Med. Sci. 58:81–83. Matar, G. M., B. Swaminathan, S. B. Hunter, L. N. Slater, and D. F. Welch. 1993. Polymerase chain reaction-based restriction fragment length polymorphism analysis of a fragment of the ribosomal operon from Rochalimaea species for subtyping. J. Clin. Microbiol. 31:1730–1734. Matteelli, A., F. Castelli, A. Spinetti, F. Bonetti, S. Graifenberghi, and G. Carosi. 1994. Short report: Verruga peruana in an Italian traveler from Peru. Am. J. Trop Med. Hyg. 50:143–144. Maurin, M., V. Roux, A. Stein, F. Ferrier, R. Viraben, and D. Raoult. 1994. Isolation and characterization by immunofluorescence, sodium dodecyl sulfate-polyacrylamide gel electrophoresis, western blot, restriction fragment length polymorphism-PCR, 16S rRNA gene sequencing, and pulsed-field gel electrophoresis of Rochalimaea quintana from a patient with bacillary angiomatosis. J. Clin. Microbiol. 32:1166–1171. Maurin, M., and D. Raoult. 1996. Bartonella (Rochalimaea) quintana infections. Clin. Microbiol. Rev. 9:273–292. McClure, W. R. 1985. Mechanism and control of transcription initiation in prokaryotes. Ann. Rev. Biochem. 54:171– 204. McGill, S. L., R. L. Regnery, and K. L. Karem. 1998. Characterization of human immunoglogublin (Ig) isotype and IgG subclass response to Bartonella henselae infection. Infect. Immun. 66:5915–5920. McGinnis-Hill, E., A. Raji, M. S. Valenzuela, F. Garcia, and R. Hoover. 1992. Adhesion to and invasion of cultured human cells by Bartonella bacilliformis. Infect. Immun. 60:4051–4058. McNee, J. W., and A Renshaw. 1916. “Trench fever”: A relapsing fever occurring with the British forces in France. Br. Med. J. 1:225–234. McCrary, B., W. Cockerham, and P. Pierce. 1994. Neuroretinitis in cat-scratch disease associated with the macular star. Pediatr. Infect. Dis. J. 13:838–839. Mehock, J. R., C. E. Greene, F. C. Gherardini, T. W. Hahn, and D. C. Krause. 1998. Bartonella henselae invasion of feline erythrocytes in vitro. Infect. Immun. 66:3462– 3466. Mernaugh, G., and G. M. Ihler. 1992. Deformation factor: An extracellular protein synthesized by Bartonella bacilliformis that deforms erythrocyte membranes. Infect. Immun. 60:937–943. Merrell, B. R., E. Weiss, and G. A. Dasch. 1978. Morphological and cell association characteristics of Rochalimaea quintana: Comparison of the vole and Fuller strains. J. Bacteriol. 135:633–640. Milam, M. W., M. J. Balerdi, J. F. Toney, P. R. Foulis, C. P. Milam, and R. H. Behnke. 1990. Epithelioid angiomato-

The Genus Bartonella

489

sis secondary to disseminated cat scratch disease involving the bone marrow and skin in a patient with acquired immune deficiency syndrome: A case report. Am. J. Med. 88:180–183. Milde P., M. Brunner, F. Borchard, T. Sudhoff, M. Burk, M. Zumdick, G. Goerz, and T. Ruzicka. 1995. Cutaneous bacillary angiomatosis in a patient with chronic lymphocytic leukemia. Arch. Dermatol. 131:933–936. Miller, V. L., J. B. Bliska, and S. Falkow. 1990. Nucleotide sequence of the Yersinia enterocolitica ail gene and characterization of the Ail protein product. J. Bacteriol. 172:1062–1069. Minnick, M. F. 1994. Identification of outer membrane proteins of Bartonella bacilliformis. Infect. Immun. 62:2644–2648. Minnick, M. F., S. J. Mitchell, and S. J. McAllister. 1996. Cell entry and the pathogenesis of Bartonella infections. Trends Microbiol. 4:343–347. Minnick, M. F. 1997a. Virulence determinants of Bartonella bacilliformis. In: B. Anderson, M. Bendinelli, and H. Friedman (Eds.) Rickettsial Infection and Immunity. Plenum Press. New York , NY. 197–211. Minnick, M. F., and K. D. Barbian. 1997b. Identification of Bartonella using PCR; genus- and species-specific primer sets. J. Microbiol. Meth. 31:51–57. Mitchell, S. J., and M. F. Minnick. 1995. Characterization of a two-gene locus from Bartonella bacilliformis associated with the ability to invade human erythrocytes. Infect. Immun. 63:1552–1562. Mitchell, S. J., and M. F. Minnick. 1997a. A carboxy-terminal processing protease gene is located immediately upstream of the invasion-associated locus from Bartonella bacilliformis. Microbiol. 143:1221–1233. Mitchell, S. J., and M. F. Minnick. 1997b. Cloning, functional expression, and complementation analysis of an inorganic pyrophosphatase from Bartonella bacilliformis. Can. J. Microbiol. 43:734–743. Moulder, J. W. 1974. Order I: Rickettsiales, (Giewszczkiewicz) 1939. In: R. E. Buchanan and N. E. Gibbons (Eds.) Bergey’s Manual of Determinative Bacteriology, 8th ed. Williams and Wilkins. Baltimore, MD. 25:687. Murakawa, G. J. 1997. Pathogenesis of Bartonella henselae in cutaneous and systemic disease. J. Am. Acad. Dermatol. 37:775–776. Musso, D., M. Drancourt, and D. Raoult. 1995. Lack of bactericidal effect of antibiotics except aminoglycosides on Bartonella (Rochalimaea) henselae. J. Antimicrob. Agents Chemother. 36:101–108. Myers,W. F., L. D. Cutler, and C. L. Wisseman. 1969. Role of erythrocytes and serum in the nutrition of Rickettsia quintana. J. Bacteriol. 97:663–666. Myers, W. F., and C. L. Wisseman. 1978. Effect of specific antibody and complement on the survival of Rochalimaea quintana in vitro. Infect. Immun. 22:288–289. Myers, W. F., C. L. Wisseman, P. Fiset, E. V. Oaks, and J. F. Smith. 1979. Taxonomic relationship of vole agent to Rochalimaea quintana. Infect. Immun. 26:976–983. Norman, A. F., R. Regnery, P. Jameson, C. Greene, and D. C. Krause. 1995. Differentiation of Bartonella-like isolates at the species level by PCR-restriction fragment length polymorphism in the citrate synthase gene. J. Clin. Microbiol. 33:1797–1803. O’Connor, S. P., M. Dorsch, A. G. Steigerwalt, D. J. Brenner, and E. Stackebrandt. 1991. 16S rRNA sequences of Bartonella bacilliformis and cat scratch disease bacillus

490

M.F. Minnick and B.E. Anderson

reveal phylogenetic relationships with the alpha-2 subgroup of the class Proteobacteria. J. Clin. Microbiol. 29:2144–2150. Padmalayam, I., B. Anderson, M. Kron, T. Kelly, and B. Baumstark. 1997. The 75-kilodalton antigen of Bartonella bacilliformis is a structural homolog of the cell division protein FtsZ. J. Bacteriol. 179:4545–4552. Padmalayam, I., K. Karem, B. Baumstark, and R. Massung. 2000a. The gene encoding the 17-kDa antigen of Bartonella henselae is located within a cluster of genes homologous to the virB virulence operon. DNA Cell Biol. 19:377–382. Padmalayam, I., T. Kelly, B. Baumstark, and R. Massung. 2000b. Molecular cloning, sequencing, expression and characterization of an immunogenic 43-kilodalton lipoprotein of Bartonella bacilliformis that has homology to NlpD/LppB. Infect. Immun. 68:4972–4979. Patel, R., J. O. Newell, G. W. Prokopp, and D. H. Persing. 1999. Use of polymerase chain reaction for citrate synthase gene to diagnose Bartonella quintana endocarditis. Am. J. Clin. Pathol. 112:36–40. Patnaik, M., W. A. Schwartzman, N. E. Barka, and J. B. Peter. 1992. Possible role of Rochalimaea henselae in pathogenesis of AIDS encephalopathy. Lancet 340:971. Perkins, B. A., B. Swaminathan, L. A. Jackson, D. J. Brenner, J. D. Wenger, R. L. Regnery, and D. J. Wear. 1992. Case 22-1992-pathogenesis of cat scratch disease [letter]. N. Engl. J. Med. 327:1599–1600. Perkocha, L. A., S. M. Geaghan, T. S. Yen, S. L. Nishimura, S. P. Chan, R. Garcia-Kennedy, G. Honda, A. C. Stoloff, H. Z. Klein, R. L. Goldman, S. VanMeter, L. D. Ferrell, and P. E. LeBoit. 1990. Clinical and pathological features of bacillary peliosis hepatis in association with human immunodeficiency virus infection. N. Engl. J. Med. 323:1581–1586. Plotkin, G. R. 1980. Agrobacterium radiobacter prosthetic valve endocarditis. Ann. Intern. Med. 93:839–840. Raoult, D., P. E. Fournier, M. Drancourt, T. J. Marrie, J. Etienne, J. Cosserat, P. Cacoub, Y. Poinsignon, P. Leclercq, and A. M. Sefton. 1996. Diagnosis of 22 new cases of Bartonella endocarditis. Ann. Intern. Med. 125:646–652. Regnath, T., M. E. Mielke, M. Arvand, and H. Hahn. 1998. Murine model of Bartonella henselae infection in the immunocompetent host. Infect. Immun. 66:5534–5536. Regnery, R. L., C. L. Spruill, and B. D. Plikaytis. 1991. Genotypic identification of rickettsiae and estimation of intraspecies sequence divergence for portions of two rickettsial genes. J. Bacteriol. 173:1576–1589. Regnery, R. L., B. E. Anderson, J. E. Clarridge, M. C. Rodriguez-Barradas, D. C. Jones, and J. H. Carr. 1992a. Characterization of a novel Rochalimaea species, R. henselae, sp. nov., isolated from blood of a febrile, human immunodeficiency virus-positive patient. J. Clin. Microbiol. 30:265–274. Regnery, R. L., J. G. Olson, B. A. Perkins, and W. Bibb. 1992b. Serological response to “Rochalimaea henselae” antigen in suspected cat-scratch disease. Lancet 339:1443–1445. Regnery, R., and J. Tappero. 1995. Unraveling mysteries associated with cat-scratch disease, bacillary angiomatosis, and related syndromes. Emerg. Infect. Dis. 1:16–21. Reidl, J., and J. J. Mekalanos. 1996. Lipoprotein e(P4) is essential for hemin uptake by Haemophilus influenzae. J. Exp. Med. 183:621–629. Relman, D. A., J. S. Loutit, T. M. Schmidt, S. Falkow, and L. S. Tompkins. 1990. The agent of bacillary angiomato-

CHAPTER 3.1.18 sis: An approach to the identification of uncultured pathogens. N. Engl. J. Med. 323:1573–1580. Relman, D. A., P. W. Lepp, K. N. Sadler, and T. M. Schmidt. 1992. Phylogenetic relationships among the agent of bacillary angiomatosis, Bartonella bacilliformis, and other alpha proteobacteria. Molec. Microbiol. 6:1801– 1807. Resto-Ruiz, S., D. Sweger, R. H. Widen, N. Valkov, and B. E. Anderson. 2000. Transcriptional activation of the high temperature requirement A (htrA) gene from Bartonella henselae. Infect. Immun. 68:5970–5978. Reynafarje, C., and J. Ramos. 1961. The hemolytic anemia of human bartonellosis. Blood 17:562–578. Ricketts, W. E. 1949. Clinical manifestations of Carrion’s disease. Arch. Intern. Med. 84:751–781. Roberts Jr., N. J. 1995. Bartonella bacilliformis (bartonellosis). In: G. L. Mandell, J. E. Bennett, and R. Dolin (Eds.) Principles and Practice of Infectious Diseases, 4th ed. Livingstone Press. New York, NY. 2209–2210. Rodriguez-Barradas, M. C., J. C. Bandres, R. J. Hamill, J. Trial, J. E. Clarridge, R. E. Baughn, and R. D. Rossen. 1995a. In vitro evaluation of the role of humoral immunity against Bartonella henselae. Infect. Immun. 63:2367–2370. Rodriguez-Barradas, M. C., R. J. Hamill, E. D. Houston, P. R. Georghiou, J. E. Clarridge, R. L. Regnery, and J. E. Koehler. 1995b. Genomic fingerprinting of Bartonella species by repetitive-element PCR for distinguishing species and isolates. J. Clin. Microbiol. 33:1089–1093. Roux, V., and D. Raoult. 1995. Inter- and intraspecies identification of Bartonella (Rochalimaea) species. J. Clin. Microbiol. 33:1573–1579. Roux, V., S. J. Eykyn, S. Wyllie, and D. Raoult. 2000. Bartonella vinsonii subsp berkhoffii as an agent of afebrile blood culture-negative endocarditis in a human. J. Clin. Microbiol. 38:1698–1700. Sander, A. 1998. Katzenkratzkrankheit IV-1. 19. In: F. Hofmann (Ed.) Infektiologie. Ecomed Verlag. Erg.Lfg., 24. Sander, A., A. Zagrosek, W. Bredt, E. Schlitz, Y. Piemont, C. Lanz, and C. Dehio. 2000. Characterization of Bartonella clarridgeiae flagellin (FlaA) and detection of antiflagellin antibodies in patients with lymphadenopathy. J. Clin. Microbiol. 38:2943–2948. Scherer, D. C., I. DeBuron-Connors, and M. F. Minnick. 1993. Characterization of Bartonella bacilliformis flagella and effect of antiflagellin antibodies on invasion of human erythrocytes. Infect. Immun. 61:4962–4971. Schmiederer, M., and B. Anderson. 2000. Cloning, sequencing, and expression of three Bartonella henselae genes homologous to the Agrobacterium tumefaciens VirB region. DNA Cell Biol. 19:141–147. Schwartzman, W. A. 1992. Infections due to Rochalimaea: The expanding clinical spectrum. Clin. Infect. Dis. 15:893–900. Slater, L. N., D. F. Welch, D. Hensel, and D. W. Coody. 1990. A newly recognized fastidious gram-negative pathogen as a cause of fever and bacteremia. N. Engl. J. Med. 323:1587–1593. Slater, L. N., D. F. Welch, and K. Min. 1992. Rochalimaea henselae causes bacillary angiomatosis and peliosis hepatis. Arch. Intern. Med. 152:602–606. Slater, L. N., J. V. Pitha, L. Herrera, M. D. Hughson, K.-W. Min, and J. A. Reed. 1994. Rochalimaea henselae infection in acquired immunodeficiency syndrome causing inflammatory disease without angiomatosis or peliosis:

CHAPTER 3.1.18 Demonstration by immunocytochemistry and corroboration by DNA amplification. Arch. Pathol. Lab. Med. 118:33–38. Smith, D. L. 1997. Cat-scratch disease and related clinical syndromes. Am. Fam. Physician 55:1783–1794. Soheilian, M., N. Markomichelakis, and C. S. Foster. 1996. Intermediate uveitis and retinal vasculitis as manifestations of cat scratch disease. Am. J. Ophthalmol. 122:582– 584. Southern, P. M. 1996. Bacteremia due to Agrobacterium tumefaciens (radiobacter): Report of infection in a pregnant woman and her stillborn fetus. Diagn. Microbiol. Infect. Dis. 24:43–45. Spach, D. H., L. A. Panther, D. R. Thorning, J. E. Dunn, J. J. Plorde, and R. A. Miller. 1992. Intracerebral bacillary angiomatosis in a patient infected with human immunodeficiency virus. Ann. Intern Med. 116:740–742. Spach, D. H., K. P. Callis, D. S. Paauw, Y. B. Houze, F. D. Schoenknecht, D. F. Welch, H. Rosen, and D. J. Brenner. 1993. Endocarditis caused by Rochalimaea quintana in a patient infected with human immunodeficiency virus. J. Clin. Microbiol. 31:692–694. Spach, D. H., A. S. Kanter, M. J. Dougherty, A. M. Larson, M. B. Coyle, D. J. Brenner, B. Swaminathan, G. M. Matar, D. F. Welch, R. K. Root, and W. E. Stamm. 1995. Bartonella (Rochalimaea) quintana bacteremia in innercity patients with chronic alcoholism. N. Engl. J. Med. 332:424–428. Stoler, M. H., T. A. Bonfiglio, R. T. Steigbigel, and M. Pereira. 1983. An atypical subcutaneous infection associated with acquired immune deficiency syndrome. Am. J. Clin. Pathol. 80:714–718. Strong, R. P. (Ed.). 1918. Trench fever: Report of Commission, Medical Research Committee, American Red Cross. Oxford University Press. Oxford , 40–60. Sweger, D., S. Resto-Ruiz, D. P. Johnson, M. Schmiederer, N. Hawke, and B. Anderson. 2000. Conservation of the 17kilodalton antigen gene within the genus Bartonella. Clin. Diag. Lab. Immunol. 7:251–257. Tappero, J. W., J. E. Koehler, T. G. Berger, C. J. Cockerell, T.-H. Lee, M. P. Busch, D. P. Stites, J. Mohle-Boetani, A. L. Reingold, and P. E. LeBoit. 1993a. Bacillary angiomatosis and bacillary splenitis in immunocompetent adults. Ann. Intern. Med. 118:363–365. Tappero, J. W., J. Mohle-Boetani, J. E. Koehler, B. Swaminathan, T. G. Berger, P. E. LeBoit, L. L. Smith, J. D. Wenger, R. W. Pinner, C. A. Kemper, and A. L. Reingold. 1993b. The epidemiology of bacillary angiomatosis and bacillary peliosis. JAMA 269:770–775. Tsukahara, M., H. Tsuneoka, H. Lino, I. Murano, H. Takahashi, and M. Uchida. 2000. Bartonella henselae infection as a cause of fever of unknown origin. J. Clin. Microbiol. 38:1990–1991. Tyeryar, F. J., E. Weiss, D. B. Millar, F. M. Bozeman, and R. A. Ormsbee. 1973. DNA base composition of rickettsiae. Science 180:415–417. Umemori, E., Y. Sasaki, K.-I. Amano, and Y. Amano. 1992. A phage in Bartonella bacilliformis. Microbiol. Immunol. 36:731–736. Urteaga, B. O., and E. H. Payne. 1955. Treatment of the acute febrile phase of Carrion’s disease with chloramphenicol. Am. J. Trop. Med. 4:507–511. Varela, G., J. W. Vinson, and C. Molina-Pasquel. 1969. Trench fever. II: Propagation of Rickettsia quintana on cell-free medium from the blood of two patients. Am. J. Trop. Med. Hyg. 18:708–712.

The Genus Bartonella

491

Verma, A., G. E. Davis, and G. M. Ihler. 2000. Infection of human endothelial cells with Bartonella bacilliformis is dependent on rho and results in activation of rho. Infect. Immun. 68:5960–5969. Vinson, J. W., and H. S. Fuller. 1961. Studies on trench fever. I: Propagation of rickettsia-like microorganisms from a patient’s blood. Path. Microbiol. 24:152–166. Vinson, J. W., G. Varela, and C. Molina-Pasquel. 1969. Trench fever. III: Induction of clinical disease in volunteers inoculated with Rickettsia quintana propagated on blood agar. Am. J. Trop. Med. Hyg. 18:713–722. Waldvogel, K., R. L. Regnery, B. E. Anderson, R. Caduff, J. Caduff, and D. Nadal. 1994. Disseminated cat-scratch disease: Detection of Rochalimaea henselae in affected tissue. Eur. J. Pediatr. 153:23–27. Walker, T. S., and H. H. Winkler. 1981. Bartonella bacilliformis: Colonial types and eyrthrocyte adherence. Infect. Immun. 31:480–486. Webster, G. F., C. J. Cockerell, and A. E. Friedman-Kien. 1992. The clinical spectrum of bacillary angiomatosis. Br. J. Dermatol. 126:535–541. Weinman, D. 1965. The bartonella group. In: R. J. Dubos and J. G. Hirsch (Eds.) Bacterial and Mycotic Infections of Man. Lippincott. Philadelphia, PA. 775–785. Weinman, D. 1974. Family II: Bartonellaceae, (Giewszczkiewicz) 1939. In: R. E. Buchanan and N. E. Gibbons (Eds.) Bergey’s Manual of Determinative Bacteriology, 8th ed. Williams and Wilkins. Baltimore, MD. 25:717– 719. Weiss, E., and G. A. Dasch. 1982. Differential characteristics of strains of Rochalimaea: Rochalimaea vinsonii sp. nov., the Canadian vole agent. Int. J. Syst. Bacteriol. 32:305–314. Weiss, E., and J. W. Moulder. 1984. Order I: Rickettsiales. In: N. R. Krieg (Ed.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 687– 719. Welch, D. F., D. A. Pickett, L. N. Slater, A. G. Steigerwalt, and D. J. Brenner. 1992. Rochalimaea henselae, sp. nov., a cause of septicemia, bacillary angiomatosis, and parenchymal bacillary peliosis. J. Clin. Microbiol. 30:275–280. Welch, D. F., D. M. Hensel, D. A. Pickett, V. H. San-Joaquin, A. Robinson, and L. N. Slater. 1993. Bacteremia due to Rochalimaea henselae in a child: Practical identification of isolates in the clinical laboratory. J.Clin. Microbiol. 31:2381–2386. Welch, D. F., K. C. Carroll, E. K. Hofmeister, D. H. Persing, D. A. Robinson, A. G. Steigerwalt, and D. J. Brenner. 1999. Isolation of a new subspecies, Bartonella vinsonii subsp. arupensis, from a cattle rancher: Identity with isolates found in conjuction with Borrelia burgdorferi and Babesia microti among naturally infected mice. J. Clin. Microbiol. 37:2598–2601. Westfall, H. N., D. C. Edman, and E. Weiss. 1984. Analysis of fatty acids of the genus Rochalimaea by electron capture gas chromatography: Detection of nonanoic acid. J. Clin Microbiol. 19:305–310. Wilfert, C. M. 1992. Brucella. In: W. K. Joklik, H. P. Willett, D. B. Amos, and C. M. Wilfert (Eds.) Zinsser Microbiology. Appleton and Lange. Norwalk, CT. 609–614. Williams-Bouyer, N. M., and E. M. Hill. 1999. Involvement of host cell tyrosine phosphorylation in the invasion of Hep-2 cells by Bartonella bacilliformis. FEMS Microbiol. Lett. 171:191–201.

492

M.F. Minnick and B.E. Anderson

Xu, Y.-H., Z.-Y. Lu, and G. M. Ihler. 1995. Purification of deformin, an extracellular protein synthesized by Bartonella bacilliformis which causes deformation of erythrocyte membranes. Biochim. Biophys. Acta 1234:173–183.

CHAPTER 3.1.18 Zangwill, K. M., D. H. Hamilton, B. A. Perkins, R. L. Regnery, B. D. Plikaytis, J. L. Hadler, M. L. Cartter, and J. D. Wenger. 1993. Cat scratch disease in Connecticut: Epidemiology, risk factors, and evaluation of a new diagnostic test. N. Engl. J. Med. 329:8–13.

Prokaryotes (2006) 5:493–528 DOI: 10.1007/0-387-30745-1_20

CHAPTER 3.1.19 ehT

redrO

se l a i s t t ekc iR

The Order Rickettsiales XUE-JIE YU AND DAVID H. WALKER

Introduction Rickettsiae are genetically related α-Proteobacteria with fascinating obligately intracellular lifestyles usually involving alternation between vertebrate and invertebrate hosts. These feats are accomplished with small, evolutionarily selected genomes. The agents classified as Rickettsia, Orientia, Ehrlichia, Anaplasma, Wolbachia and Neorickettsia cause diseases (such as epidemic louse-borne typhus fever, Rocky Mountain spotted fever, scrub typhus, human ehrlichioses, and bovine anaplasmosis) that have repeatedly altered the course of history (Zinsser, 1935), unexpectedly kill previously healthy tickexposed persons, occur as highly prevalent endemic febrile illness, and are veterinary pathogens of major economic importance. Although mostly known as pathogens transmitted by hematophagous ticks and insects, some of the Rickettsiales are insect endosymbionts (Wolbachia) that have evolved dramatic abilities to manipulate their host populations, and others (Neorickettsia) have a life cycle within a trematode that involves a series of aquatic hosts including snails, fish and insects. The biology of the rickettsiae is difficult to define and investigate separately from the biology of their hosts. As a result of obligately intracellular growth and propagation difficulties in antibiotic-free cell culture or even in embryonated eggs, rickettsial physiology and metabolism are challenges to elucidate. Consequently, the study of these microorganisms and their diseases has been relatively neglected and there is the mistaken impression that they must be unimportant and uninteresting. Nothing could be further from the truth. The state of Rickettsiales knowledge presented in this chapter is to suggest to microbiologists opportunities for increasing scientific and medical understanding in this field. On the basis of phylogenetic analysis of the 16S rRNA gene sequence (Fig. 1), the order Rickettsiales is a monophyletic group (Roux and Raoult, 1999) that currently includes the families

Rickettsiaceae and Anaplasmataceae (Stothard and Fuerst, 1995; Dumler et al., 2001). All bacteria in the order Rickettsiales are Gramnegative α-Proteobacteria. The arrangement of the rRNA genes in Rickettsiales is unusual. While the 16S, 23S, and 5S rRNA genes are linked together in other bacteria, in rickettsial organisms, the 16S rRNA gene is separated from the 23S and 5S rRNA gene cluster, and the 23S rRNA gene is preceded by a gene that codes for methionyl-tRNA formyltransferase (Andersson et al., 1999; Massung et al., 2001). The genomic rearrangement of the rRNA genes preceded the divergence of the two families. On the basis of phylogenetic analysis of 16S rRNA gene sequences, many species classified traditionally as Rickettsiales do not cluster with and have been removed from the Rickettsiaceae and Anaplasmataceae families. These include Rickettsiella grylli, Coxiella burnetii, Wolbachia persica, Bartonella, Grahamella, Eperythrozoon ovis, Hemobartonella felis and H. muris (Roux and Raoult, 1999). The family Anaplasmataceae includes four genera, Anaplasma, Ehrlichia, Neorickettsia and Wolbachia. Members of the genera Anaplasma, Ehrlichia, and Neorickettsia in the family Anaplasmataceae are small obligately intracellular, Gram-negative, pleomorphic, coccoid to ellipsoidal organisms that reside in membranebound cytoplasmic vacuoles and form characteristic microcolonies resembling mulberries, termed morulae (Latin morum = mulberry; Fig. 2). Electron microscopy reveals two distinct morphological forms: a larger reticulate and a smaller dense-core cell. Both forms divide by binary fission, which is strong evidence that they are not stages in a developmental cycle (Popov et al., 1998; Popov et al., 2000). All known members of these genera are pathogenic for mammals, and some are pathogenic for human beings. Most members of the Anaplasmataceae target hematopoietic cells. Anaplasma marginale infects the erythrocytes of cattle and wild ruminants, and A. bovis infects the mononuclear cells of cattle. Anaplasma phagocytophila and

494

X.-J. Yu and D.H. Walker

CHAPTER 3.1.19

100

R. honei R. africae R. sikirica R. conorii R. parkeri R. prickettsii R. skwaca

100

R. japonica 88 HLJ054 R.amblyommii

10 changes 100

R. asschlimamii R. rhipicephali R. massiliae

Rickettsia

R. akari 100

100

100

100

100

75

R. promazekii R. typhi R. hetretica 50 R. australis 100 R. bellii R. canaclensis 100 R. montanensis O. tsutsugamushi 51 E. chatieensis 51 E. ewingii 87 E. canis 100 E. muris E. ruminantium 72 A. platges A. phagocytophila 100 A. ovis 100 60 A. marginale A. centrale 100 A. bovis W. pipientis

100

63

100

Orientia

Ehrilchia

Anaplasma

Wolbachia

N. risticii N. sennetsu

Neorickettsia

SF agent N. helminthoeca Fig. 1. Phylogenetic relationships of the organisms in the order Rickettsiales based on the DNA sequences of the 16S rRNA genes (GenBank accession numbers: R. honei, AF060705; R. africae, RIRRGDA; R. sibirica, RIRRS16SRG; R. conorii, RIRRGDH; R. parkeri, RIRRRDA; R. rickettsii, RIRRGDP; R. slovaca, RIRRGDX; R. japonica, RIRRGDL; Strain HLJ054, AF178037; R. amblyommii, RAU11012; R. aeschlimannii, RAU74757; R. rhipicephali, RIRRGDO; R. massiliae, RIRRGDI; R. akari, RAU12458; R. prowazekii, RIRGGSA; R. typhi, RIRRGDU; R. helvetica, RIRRGDK; R. australis, RAU17644; R. bellii, RBU11014; R. canadensis, RCU15162; R. montanensis, RIRRGDN; O. tsutsugamushi, RIRRTKP16B; E. chaffeensis, AF147752; E. ewingii, EEU96436; E. canis, AF162860; E. muris, EMU15527; E. ruminantium, CRDNA; A. platys, AF156784; A. phagocytophila, AY055469.1; A. ovis, AF309865; A. marginale, APMRR16SA; A. centrale, AF318944; A. bovis, EBU03775; W. pipientis, U23709; N. risticii, EHRRGBSA; N. sennetsu, EHRRRNAI; SF agent, EHRSF; and N. helminthoeca, NHU12457). The length of each pair of branches represents the distance between sequence pairs. The numbers on the branch indicate the bootstrap values.

A. platys infect mammalian neutrophils and canine platelets, respectively. All members of the genus Ehrlichia except E. ruminantium and E. ewingii infect monocytes. Ehrlichia ruminantium infects endothelium, and E. ewingii infects granulocytes.

Wolbachia pipientis and the genus Anaplasma form sister taxons by 16S rRNA gene and groEL gene sequence phylogeny (Wen et al., 1995; Yu et al., 2001). Thus, W. pipientis was reclassified into the family Anaplasmataceae (Dumler et al., 2001).

CHAPTER 3.1.19

The Order Rickettsiales

495

R. honei 1 um R. peacockii 72 62 5 changes

R. rickettsi R. parkeri

89

R. conorii R. japonica

71 R. montanensis R. rhipicephali 100 R. amblyommii 57 Fig. 2. Electron photomicrograph of a canine macrophagelike cell line (DH82) infected with Ehrlichia canis contains eight vacuoles (morulae) filled with ehrlichiae. Five morulae contain reticulate cell forms; three contain dense-core forms (particularly the morula in the lower center of the figure). See the Microbe Library website (http://www.microbelibrary. org/) for more electron micrographs of Ehrlichia.

Genus Rickettsia Phylogeny The evolution of the genus Rickettsia is demonstrated by phylogenetic trees developed from DNA sequence data that are based on the 16S rRNA, groEL, citrate synthase (glt), rompA, rompB and other genes (Roux and Raoult, 1995; Stothard and Fuerst, 1995; Roux et al., 1997; Fournier et al., 1998; Roux and Raoult, 2000; Fig. 3). The most ancestral species of the genus appear to be R. bellii, R. canadensis, and the AB male killing bacterium (Stothard et al., 1994). Rickettsia and mitochondria evolved from a common ancestor (Andersson et al., 1998). The phylogenetic tree may be overpopulated by species’ names of spotted fever group rickettsiae, many of which are closely related and have been given separate names. The discovery of organisms very closely related to R. bellii in herbivorous pea aphids and associated with plant pathology (e.g., papaya bunchy top disease) suggests that knowledge and diversity of Rickettsia may have been determined more by medical investigations and medical entomology than by consideration of the complete picture (Chen et al., 1996; Davis et al., 1998).

R. felis R. australis R. helvetica

R. typhi R. prowazelai Fig. 3. Phylogenetic relationships of the organisms in the genus Rickettsia based on the DNA sequences of the 17-kDa protein genes (GenBank accession numbers for R. honei, A060704; R. peacockii, AF260571; R. rickettsii, RIRANTRR; R. parkeri, RPU17008; R. conorii, RIRANT17KA; R. japonica, RIR17KGCA; R. montanensis, RMU11017; R. rhipicephali, RPU11020; R. amblyommii, RAU11013; R. felis AF195118; R. australis, RIRTRAPRO; and R. helvetica, AF181036). The length of each pair of branches represents the distance between sequence pairs. The numbers on the branch indicate the bootstrap values.

Taxonomy The family Rickettsiaceae consists of two genera: Rickettsia and Orientia. Rickettsia is further divided into two groups: the spotted fever group (SFG) and typhus group (TG), based on the difference in lipopolysaccharide (LPS) antigens. The TG rickettsiae include R. prowazekii and R. typhi. The SFG rickettsiae include R. akari, R. australis, R. africae, R. conorii, R. honei, R. japonica, R. sibirica R. helvetica, R. slovaca, R. massiliae, R. rhipicephali, R. aeschlimannii, R. montanensis, R. parkeri, and most likely R. felis (Roux and Raoult, 2000; Zhang et al., 2000; Bouyer et al., 2001). The numbers of named SFG rickettsiae have increased rapidly in recent years

496

X.-J. Yu and D.H. Walker

owing to improved isolation approaches and molecular methods for identification; however, there is no consensus about the criteria used to define a species of SFG rickettsiae. Many designated species of SFG rickettsiae actually may be clones or strains of a single species, according to the standards used for classifying other bacteria. Other Rickettsia (such as R. bellii, R. canadensis and the AB male killing bacterium) do not fit into either the SFG or TG.

Habitat Rickettsia are small (0.3–0.5 × 0.8–2.0 µm), Gram-negative, aerobic coccobacilli. They are obligately intracellular and reside free in the cytoplasm of the eukaryotic host cell where they divide by binary fission. SFG rickettsiae may also reside in the nucleus of the eukaryotic host cells. Rickettsia are dependent on arthropods (ticks, mites, fleas and lice) for their persistence in nature. For some, the arthropod host is both reservoir and vector. Transovarian transmission of the agent from the infected female to the next generation through the ova is the essential mechanism for many species (Burgdorfer, 1988).

Isolation Isolation of rickettsiae from blood, buffy coat, or plasma historically employed inoculation of adult male guinea pigs or embryonated chicken eggs for Rickettsia and mice for Orientia (Walker, 1996a). More often at present Vero, L-929, HEL, and MRC5 cells in antibiotic-free media are inoculated with heparin-anticoagulated plasma or buffy coat ideally collected before antirickettsial treatment. Isolation of rickettsiae is enhanced by centrifugation of the inoculum onto monolayers in shell vials (LaScola and Raoult, 1996). Rickettsiae are detected by examining monolayers stained by Giemsa, Gimenez, or immunofluorescence methods, with 82% of positive samples identified after 48 hours incubation. Appropriate precautions should be taken for handling these P3 level pathogens.

Identification Members of the genus Rickettsia have a typical Gram-negative bacterial cell wall that contains LPS, peptidoglycan, and outer membrane proteins. Isolated organisms can be identified as Rickettsia by the obligate intracellular parasitism, ultrastructural identification of cell wall morphology and of location free in the cytosol, reactivity with group-, species-, or strain-specific monoclonal or polyclonal antibodies, and restriction fragment length polymorphism of particular

CHAPTER 3.1.19

genes. Currently DNA sequence analysis of the genes of the 17-kDa lipoprotein, outer membrane proteins A and B, citrate synthase, and 16S rRNA is utilized most often to identify rickettsial isolates (Walker, 1996a).

Preservation Stocks of Rickettsia are usually preserved frozen at –70 to –80°C or in the vapor phase of liquid nitrogen. They may also be preserved in a lyophilized state.

Physiology Adherence to the Host Cell Rickettsiae are inoculated into the dermis of the skin by a tick bite or through damaged skin from the feces of lice or fleas. Rickettsiae spread through the bloodstream and infect the endothelium. Adherence to the host cell is the first step of rickettsial pathogenesis. The adhesins must be outer membrane proteins. The outer membrane protein OmpA has been implicated as an adhesin of R. rickettsii because monoclonal antibodies to OmpA block R. rickettsii attachment (Li and Walker, 1998). The rickettsial adhesin must have evolved prior to the divergence of the SFG and TG rickettsiae because both groups are obligately intracellular. Lack of OmpA in TG rickettsiae indicates that there must be a more conserved or even more important adhesin in rickettsiae than OmpA for mammalian cells. There is evidence that both OmpA and OmpB are adhesins of R. japonica (Uchiyama, 1999). The receptor for Rickettsia has yet to be identified. Although the main target cells of Rickettsia in vivo are endothelial cells, rickettsiae can infect virtually every cell line in vitro. Thus, either the receptor for Rickettsia is ubiquitous among cells, or rickettsiae can bind to different receptors. Invasion of the Host Cell Upon attaching to the host cell membrane, rickettsiae are phagocytosed by the host cell. Rickettsia and Orientia are believed to induce host cell phagocytosis because they can enter cells that normally do not phagocytose particles (Walker, 1984a). Once phagocytosed by the host cell, rickettsiae and orientiae are observed to enter quickly the cytoplasm outside of the phagosome. Thus, they are said to escape the phagosome. Neither the molecules nor the mechanisms involved in rickettsial escape from the phagosome have been determined. Phospholipase A2 (PLA2) has been suggested to be involved in the lysis of the phagosomal membrane by Rickettsia (Winkler and Miller, 1982; Walker et al., 1984b; Winkler

CHAPTER 3.1.19

and Daugherty, 1989; Silverman et al., 1992; Manor et al., 1994; Winkler et al., 1994; Ojcius et al., 1995); however, genome sequencing did not reveal a gene encoding a PLA2. To search for the rickettsial PLA2 gene by comparing rickettsial genes with PLA2 genes of bees or snakes might not be warranted since the rickettsial PLA2 involved in the lysis of host cell membrane may be very different. We have found a rickettsial protein (Accession no. RPXX03) with a calciumindependent PLA2 motif (Yu et al., 2000c). The rickettsial protein is homologous to the exoenzyme (ExoU) of Pseudomonas aeruginosa. ExoU expression in P. aeruginosa is linked to acute cytotoxicity (Vallis et al., 1999). The enzymatic activity of the rickettsial protein has yet to be characterized. Movement within and Release from the Host Cell Observations in cell culture systems suggest that the mechanisms of intracellular movement and destruction of the host cell differ among the spotted fever and typhus group rickettsiae (Silverman and Santucci, 1988; Silverman, 1997). TG rickettsiae are released from host cells by lysis of the cells. After infection with R. prowazekii or R. typhi, the rickettsiae continue to multiply until the cell is packed with organisms and then bursts. Cell death possibly results from apoptosis or membranolytic activity that previously has been hypothesized to be due to phospholipase A2 (Winkler and Miller, 1982; Walker et al., 1984b; Winkler and Daugherty, 1989; Silverman et al., 1992; Manor et al., 1994; Winkler et al., 1994; Ojcius et al., 1995). Before lysis, TG rickettsia-infected host cells have a normal ultrastructural appearance. SFG rickettsiae seldom accumulate in large numbers and do not burst the host cells. They escape from the cell by stimulating polymerization of host cell-derived F-actin tails (Fig. 4),

1 um

Fig. 4. Electron photomicrograph of an organism of Rickettsia conorii (right) with a long actin tail extending from its left pole to the left of the figure. Photomicrograph provided by Vesevolod Popov.

The Order Rickettsiales

497

which propel them through the cytoplasm and into tips of filopodia, from which they emerge (Schaechter et al., 1957; Teysseire et al., 1992; Heinzen et al., 1993; Heinzen et al., 1999). Infected cells exhibit signs of membrane damage associated with an influx of water, which is sequestered within cisternae of dilated, rough endoplasmic recticulum (Walker and Cain, 1980a). The means by which rickettsiae damage host cell membranes is uncertain, but there is experimental evidence to suggest a role for free radicals of oxygen, a phospholipase, or a protease (Walker et al., 1983b; Walker et al., 1984b; Walker et al., 2002; Silverman and Santucci, 1988; Silverman and Santucci, 1990; Eremeeva and Silverman, 1998; Eremeeva et al., 2001). The protein responsible for the actin-based movement in SFG rickettsiae has yet to be identified. The genome sequence of R. conorii did not reveal a rickettsial protein homologous to ActA or Ics, the proteins responsible for actin polymerization by Listeria monocytogenes and Shigella flexneri, respectively; however, a hypothetical protein of 520 residues (RC0909) exhibits an overall organization similar to that of ActA. Both proteins are highly charged at the Nterminus and have a central proline-rich region. RC0909 has a weak similarity to the Wiskott-Aldrich. Syndrome protein (WASp) homology domain 2 regulates the formation of the actin filaments (Ogata et al., 2001). Reactivation of Rickettsia Rickettsia rickettsii loses its pathogenicity and virulence for guinea pigs in starved ticks (Spencer and Parker, 1923). Injection of triturated, starved, infected ticks into guinea pigs does not cause disease, but causes asymptomatic seroconversion; however, incubation of the tick vector at 37°C for 24–48 h or feeding the ticks on an animal for 10 h or longer before trituration results in clinically manifest disease in the inoculated guinea pigs. Spencer and Parker (1923) postulated that virulence of R. rickettsii in the tick vector is linked directly to the physiological state of the tick and defined this phenomenon as “reactivation.” The mechanism of rickettsial reactivation is not understood; however, the reactivation may result from growth of rickettsiae or differential expression of rickettsial virulence factors at the elevated temperature or after stimulation by components of the blood meal. Rickettsia rickettsii increases 100-fold in the hemolymph of partially engorged ticks compared to unfed infected ticks (Wike and Burgdorfer, 1972). Differential expression of rickettsial proteins and ultrastructural changes has been confirmed by immunoblot and electron microscopy. Electron microscopy reveals

498

X.-J. Yu and D.H. Walker

that reactivated R. rickettsii in ticks incubated at 37°C or in ticks fed on animals has a discrete microcapsular layer and a discrete electronlucent slime layer outside the microcapsular layer. In starved ticks, the microcapsular layer and the slime layer of rickettsiae are inconspicuous or ragged (Hayes and Burgdorfer, 1982). Immunoblot analysis indicates that rickettsial proteins of 42, 43, 48, 75 and 100 kDa are induced in a tick cell line when shifted from 28 to 34°C (Policastro et al., 1997).

Genetics The genomes of R. prowazekii and R. conorii have been completely sequenced (Andersson et al., 1998; Ogata et al., 2001). The R. conorii genome (1,268,755 bp) is slightly larger than that of R. prowazekii (1,111,523 bp). There are 804 common open reading frames (ORFs) between R. conorii, which has 1374 ORFs, and R. prowazekii, which has 834 ORFs; 552 ORFs were found only in R. conorii and 229 of them have a homologous noncoding remnant in R. prowazekii. The G+C content is 32.4 mol% for R. conorii and 29.0 mol% for R. prowazekii. The R. conorii genome contains more repetitive DNA than that of R. prowazekii. The higher G+C content of R. conorii is accounted for by the repetitive DNA, which is G+C rich (40%; Ogata et al., 2001). Rickettsia genomes are very small, only about a quarter of the genome size of Escherichia coli strain K12 (4,639,221 bp, accession no. NC000913), a free-living Gram-negative bacterium. The small genomes of Rickettsia are the result of genome reduction, which shapes the relationship of intracellular bacteria and their hosts. Many genes involved in the biosynthesis and regulation of biosynthesis of amino acids and nucleosides were deleted inasmuch as host genes fulfill these functions. Rickettsia not only contain fewer genes but also contain more noncoding sequences (19% in R. conorii and 24% in R. prowazekii) than their free-living bacterial cousins contain (Andersson et al., 1998; Ogata et al., 2001). A large portion of the non-coding DNA in rickettsiae is hypothesized to be degraded remnants of unnecessary genes that have not yet been removed from the genome (Andersson et al., 1998). Evolutionarily, rickettsiae were derived from a free-living bacterial ancestor. Once a freeliving aerobic bacterium entered and established an intracellular parasitic relationship with the pro-eukaryote cell, many chemical substrates could be readily obtained from the host cells and used without further metabolic modification by the bacteria. The genes of the bacterium involved in metabolic pathways such as

CHAPTER 3.1.19

glycolysis, fermentation and biosynthesis of small molecules were functionally similar to their host cell’s genes. These redundant genes then could mutate and become genes with new functions or pseudogenes. Pseudogenes eventually are deleted from the genome to increase the energy use efficiency of the bacterium. Owing to loss of genes essential to a free-living mode, the bacterium became further dependent on the host cell and eventually lost its ability to live outside the host cell. All the enzyme genes for the tricarboxylic acid cycle are present in the rickettsial genome. Rickettsia retains genes for ATP synthesis complexes, despite the acquisition of an ATP/ADP translocator gene (Andersson et al., 1998). The ATP/ ADP translocases are unique to Rickettsia and Chlamydia among bacteria and might have originated from plants (Wolf et al., 1999). This enzyme allows the importation of ATP from the host cell cytoplasm. The structural differences between the ATP/ADP translocases of mitochondria and those of R. prowazekii indicate their evolutionary origins are different, despite the evidence that mitochondria and rickettsiae evolved from a common ancestor (Andersson et al., 1998). Genetic Manipulation of the Rickettsial Genome No resident plasmid or bacteriophage has been found in rickettsiae (Andersson et al., 1998; Ogata et al., 2001). Lack of genetic tools to manipulate the rickettsial genome has hampered our current understanding of the biology of rickettsiae. Recent advances in transforming rickettsiae may facilitate study of their molecular biology (Troyer et al., 1999; Rachek et al., 2000). Autotransporter Proteins In Gram-negative bacteria, proteins are transported across the cytoplasmic or inner membrane and the outer membrane by several pathways including types I, II, III, IV and autotransporter secretion. Types I, II, and III secretion systems require the assistance of a variety of accessory proteins that function in combination with the general secretory pathway (Salmond and Reeves, 1993; Galan and Collmer, 1999). In contrast, autotransporters do not need accessory proteins for assistance in self-translocation. An autotransporter protein consists of three functional domains: the aminoterminal leader sequence, the secreted mature (α) protein and a carboxy-terminal (β) domain. The signal peptide inserts itself into the inner membrane and directs the export of the precursor molecule via the general secretion pathway into the periplasm. The signal peptidase cleaves the precursor, releasing the mature polypeptide into the periplasm. Once in the periplasm, the β-

CHAPTER 3.1.19

domain of the protein is inserted into the outer membrane to form a β-barrel pore, and the passenger domain is translocated to the cell surface through the pore. The β-domain of autotransporters is conserved among even distantly related bacteria, whereas passenger domains are diverse, suggesting that autotransporters were either derived originally from a single gene or from recombination of genes for unrelated proteins with the β-domain gene (Henderson et al., 1998). A striking feature of rickettsiae is that all the identified rickettsial outer membrane proteins belong to the autotransporter family. These proteins include OmpA, OmpB, and four ORFs encoding the hypothetical proteins Sca1 (accession no. PR018), Sca2 (RP081), Sca3 (RP451), and Sca5 (RP704; Andersson et al., 1998; Bouyer et al., 2001). Sca1 and Sca2 consist of only the β-domain without the passenger peptide. Thus, Sca1 and Sca2 may be involved in the transport of other rickettsial proteins. The precursor of OmpB is a 168-kDa protein that is posttranslationally processed to the 135-kDa mature OmpB by cleavage of a 32-kDa β-peptide (Hackstadt et al., 1992). The amino acid sequences of the β-peptides of all five rickettsial autotransporters are highly homologous and also share homology with autotransporters from distantly related bacteria (Henderson et al., 1998). Most of the autotransporter proteins are adhesins or proteases of bacteria, indicating that OmpA, OmpB, Sca3 and Sca5 are possibly important rickettsial virulence factors. OmpA, a 160–190-kDa protein, has been found in all tick-borne SFG but not TG rickettsiae. The flea-borne SFG rickettsia, R. felis, has an OmpA that is truncated by the presence of premature stop codons. Over 40% of the amino acid sequence of OmpA is devoted to a hydrophilic region of tandem repeat units (Anderson et al., 1990b). The repeat units are not identical and thus are divided into two types (I with 75 amino acids and II with 72 amino acids). The type II repeats are less conserved and further divided into two subtypes (IIa and IIb; Anderson et al., 1990b). The rompA sequences upstream and downstream of the repeat region are conserved among SFG rickettsial species. Although conserved among R. rickettsii, R. conorii and R. akari, the repeat sequences vary in number and order of repeat unit arrangement among SFG rickettsiae (Gilmore, 1993); however, the R. australis repeat unit differs greatly from other SFG rickettsial repeat units. Rickettsia australis has only one type of repeat unit with only 21% identity to the type I unit of R. rickettsii (Stenos and Walker, 2000). The antigenic diversity of SFG rickettsiae is determined in large part by the number, order, and type of

The Order Rickettsiales

499

repeat units. Recombinant OmpA stimulates immune protection in guinea pigs against rickettsial challenge (Sumner et al., 1995). OmpB, found in all rickettsiae, is the most abundant rickettsial surface (S-layer) protein (Ching et al., 1990) and contains species-, group- and genusspecific epitopes (Anacker et al., 1987). OmpB stimulates immune protection in mice and guinea pigs against challenge with R. typhi (Dasch et al., 1999). Type IV Secretion System Genome sequencing revealed that R. prowazekii contains the genes virB4, virB8, virB9, virB10, virB11 and virD4 (Andersson et al., 1998), which are homologues of the type IV secretion systems founded in Agrobacterium tumefaciens (virB operon; Stachel and Nester, 1986; Kuldau et al., 1990), Brucella abortus (virB; Sieira et al., 2000), E. coli (tra genes; Winans and Walker, 1985; Pohlman et al., 1994), Bordetella pertussis (ptl genes; Weiss et al., 1993; Kotob et al., 1995), Legionella pneumophila (dot-icm genes and lvh genes; Brand et al., 1994; Segal and Shuman, 1997; Segal et al., 1999), Helicobacter pylori (cag genes; Covacci et al., 1999), E. chaffeensis, A. phagocytophilum (Ohashi et al., 2002), and Wolbachia (Brand et al., 1994; Masui et al., 2000; Sieira et al., 2000). The type IV secretion systems inject either bacterial DNA or proteins directly into the cytosol of eukaryotic host cells (Winans and Walker, 1985; Weiss et al., 1993; Christie, 1997; Covacci et al., 1999). In the facultative intracellular bacteria L. pneumophila and B. abortus, the type IV secretion systems are essential for the intracellular multiplication of the two organisms (Brand et al., 1994; Sieira et al., 2000). Legionella pneumophila icm mutant is deficient in its ability to cause lysis of the host cell. The dot/icm protein secretion apparatus of L. pneumophila secretes RalF, a protein required for recruiting ADP ribosylation factor (ARF) to the phagosome containing L. pneumophila and for vacuole biogenesis (Nagai et al., 2002). ARF is a highly conserved small GTP-binding protein that acts as a key regulator of vesicle traffic from the ER and Golgi apparatus. An R. prowazekii gene (G71694) similar to the ralf gene of L. pneumophila has been found by BLASTN search (Nagai et al., 2002). Because they are homologous to the eukaryotic ARF-specific guanine nucleotide exchange factors and not found in other bacteria, RalF and its rickettsial analogue might have been acquired from a eukaryotic gene by horizontal gene transfer (Nagai et al., 2002). On the basis of similarity with the RalF of L. pneumophila, the rickettsial protein encoded by gene G71694 may be the component that is transported by the rickettsial type IV secretion system; however, unlike L. pneumophila, Rickettsia do not live in vacuoles.

500

X.-J. Yu and D.H. Walker

Thus, the function of rickettsial gene G71694 is still not clear. The role of the type IV secretion system in the pathogenesis of other bacterial infections suggests that the rickettsial virB genes may be associated with rickettsial intracellular survival.

CHAPTER 3.1.19

papaya bunchy top disease, suggests that our knowledge of the diversity and evolutionary origin of Rickettsia might be dreadfully skewed (Chen et al., 1996; Davis et al., 1998).

Epidemiology Rickettsial Lipoprotein A 17-kDa protein has been identified in all Rickettsia species (Anderson, 1990a). The corresponding gene sequence is highly conserved among rickettsial species, indicating the importance of the protein to the survival of the rickettsiae. The protein, predicted to be a lipoprotein and known to be partly surface exposed, has been speculated to play a scaffolding and protective role in the rickettsiae (Anderson, 1990a).

Ecology The importance of transovarial maintenance for the ecology of rickettsiae varies from essential (in the nonpathogenic R. peacockii where it is the only mechanism ensuring persistence in the environment) to necessary, but not sufficient (in the highly pathogenic R. rickettsii; McDade and Newhouse, 1986; Niebylski et al., 1996; Niebylski et al., 1997). The nonpathogenic organisms appear to coexist with their arthropod host, neither harming them nor possessing the virulence traits for horizontal transmission. In contrast, R. rickettsii under some circumstances harms its tick host resulting in its death or reduced production of offspring (Niebylski et al., 1999); however, the virulence mechanisms that harm the tick also permit horizontal transmission to small rodents and sufficient levels of rickettsemia to establish new lines of transmission to feeding ticks (Gage et al., 1990). In the United States, generally fewer than 5% of vector ticks are infected with rickettsiae, most of which are nonpathogenic R. montanenisis or R. bellii (Philip and Casper, 1981a; Philip et al., 1981b). Fewer than 1 in 1000 vector ticks carry virulent R. rickettsii, presumably a result of their deleterious interaction. Higher rates of rickettsial carriage by ticks are observed with less virulent pathogens. Application of specific diagnostic methods might reveal that some organisms previously of undetermined pathogenicity, such as R. amblyommii, R. rhipicephali and R. helvetica, are actually at least mildly pathogenic. This situation has occurred for R. slovaca and strain Thai tick-118 (now established as R. honei; Stenos et al., 1998). Our view of Rickettsia ecology is highly anthropocentric and hematophagous arthropodand medically oriented. Identification of strains closely related to R. bellii in herbivorous insects and even in association with a plant disease,

Twelve species of Rickettsia have been associated with human disease: R. rickettsii (Rocky Mountain spotted fever), R. conorii (boutonneuse fever, Mediterranean spotted fever, Kenyan tick typhus, Israeli spotted fever, Indian tick typhus, and Astrakhan spotted fever), R. africae (African tick-bite fever), R. sibirica (North Asian tick typhus), R. japonica (Japanese spotted fever), R. australis (Queensland tick typhus), R. honei (Flinders Island spotted fever), R. akari (rickettsialpox), R. felis (cat flea typhus), R. slovaca (tick-borne lymphadenopathy), R. prowazekii (epidemic louse-borne typhus, recrudescent typhus fever, and sylvatic typhus), and R. typhi (murine typhus). Each rickettsiosis has a particular geographic distribution determined by its natural history involving an arthropod host and in some cases a zoonotic vertebrate host (Table 1). Often during a particular season of the year, rickettsial diseases occur when humans encounter infected ticks, fleas, or mites that are seeking a blood meal. Louse-borne typhus fever occurs particularly in cold climates among impoverished persons who are infested with lice (Wolbach et al., 1922; Perine et al., 1992). Epidemics have been associated with war and natural disasters where there is crowding, prevalence of human body lice, and lack of opportunity to bathe and wash clothes (Patterson, 1993; Raoult et al., 1998).

Disease General Considerations Rickettsioses vary in clinical severity according to the virulence of the Rickettsia and host factors, such as older age, male gender, and alcoholism, and other underlying diseases (McDade, 1990; Walker, 1990). The most virulent rickettsiae are R. rickettsii and R. prowazekii, which kill a significant portion of previously healthy infected persons, unless they are treated sufficiently early in the course of illness with an effective antirickettsial agent, usually doxycycline. Rickettsia conorii and R. typhi also possess the virulence to kill 1–2% of infected persons (Ruiz-Beltran et al., 1985; Raoult et al., 1986; Dumler et al., 1991). The other spotted fever rickettsioses can cause disseminated infection with multisystem involvement and illness of two weeks or more duration, but fatalities are rare and (for some entities) have never been reported.

CHAPTER 3.1.19

The Order Rickettsiales

501

Table 1. Geographic distribution, maintenance in nature, and human transmission. Geographic distribution

Maintenance in nature

R. rickettsii

Agent

North, Central and South America

R. conorii

Mediterranean basin, Africa, Asia Sub-Saharan Africa, Caribbean Russia, China, Mongolia, Pakistan, Kazakhstan, Kirgiziya, Tadzhikistan

Transovarial maintenance in Dermacentor, Rhipicephalus, and Amblyomma ticks: less extensive horizontal transmission from tick to mammal to tick Transovarial maintenance in Rhipicephalus ticks Transovarial maintenance in Amblyomma ticks Transovarial maintenance in Dermacentor, Haemaphysalis, and Hyalomma ticks; horizontal transmission from tick to mammal to tick Presumably a transovarial tick host Transovarial transmission in Ixodes ticks Transovarial transmission in Aponomma ticks

R. africae R. sibirica

R. japonica R. australis R. honei

R. prowazekii

Japan, China Eastern Australia Southern Australian islands, Thailand United States, Ukraine, Croatia, possibly worldwide North and South America and Europe South and Central America, Africa, Asia, Mexico United States

R. prowazekii

Worldwide

R. typhi

Worldwide, predominantly tropical and subtropical

R. akari

R. felis R. prowazekii

Human transmission Tick bite

Tick bite Tick bite Tick bite

Tick bite Tick bite Tick bite

Transovarial transmission in Liponyssoides sanguineus mites; horizontal transmission from mite to mouse to mite Transovarial transmission in Ctenocephalides felis fleas Man to Pediculus humanus corporis louse to man

Mite bite

Flying squirrel to louse and flea ectoparasites to flying squirrel

Presumably fleas of flying squirrels to man None

Reactivation of latent human infection years after acute illness Rat to Xenopsylla cheopis flea to rat; opossum to cat flea C. felis to opossum

All infections with organisms of the genus Rickettsia usually begin with introduction of rickettsiae into the skin, either through a tick bite or, in the case of contaminated flea or louse feces, through cutaneous abrasions. Rickettsiae enter dermal cells including endothelium and proliferate locally intracellularly with endothelial cell-to-cell spread for most SFG rickettsioses resulting in an eschar or tache noire, a zone of dermal and epidermal necrosis approximately 1 cm in diameter with a surrounding zone of erythema (Walker et al., 1988b). Eschars do not occur in epidemic and murine typhus and are rarely observed in Rocky Mountain spotted fever and Israeli spotted fever (Walker et al., 1981; Sarov et al., 1990). SFG rickettsioses often manifest regional lymphadenopathy in the drainage of the eschar, suggesting that rickettsiae may spread via lymphatic vessels from the tick bite inoculation site early in the infection. Rickettsiae spread throughout the body and infect mainly endothelial cells, establishing many foci of contiguous infected blood vessel-lining

Unknown Louse feces scratched into skin

Flea feces scratched into skin; rubbed into conjunctiva or inhaled

cells. Injury in these local sites causes vascular damage manifesting as rash, interstitial pneumonia, encephalitis, interstitial nephritis, and interstitial myocarditis, as well as lesions in the liver, gastrointestinal wall, pancreas, or indeed nearly any vascularized tissue of the body (Walker, 1988a; Walker, 1996b). The most important pathophysiologic effect is increased vascular permeability with consequent edema, loss of blood volume, hypoalbuminemia, decreased osmotic pressure, and hypotension (Harrell and Aikawa, 1949). These effects can be life threatening in the lungs as non-cardiogenic pulmonary edema and adult respiratory distress syndrome (Donohue, 1978; Lankford and Glauser, 1980; Walker et al., 1980b). Hypovolemia and hypotension may result in prerenal azotemia or, if shock occurs, in acute tubular necrosis (Walker and Mattern, 1979). In the most severe cases of Rocky Mountain spotted fever, louse-borne typhus, boutonneuse fever, and murine typhus, central nervous system involvement is a critical life-threatening factor (Helmick

502

X.-J. Yu and D.H. Walker

et al., 1984). The precise relative contributions of the altered blood-brain barrier, hypoperfusion, hypoxemia, cerebral edema, and other pathophysiologic effects have not been determined, but progressively severe illness involves confusion, delirium, stupor, ataxia, coma, focal neurologic signs, and seizures (Miller and Price, 1972a; Miller and Price, 1972b; Horney and Walker, 1988). Cerebrospinal fluid abnormalities may include pleocytosis and increased protein concentration. Hyponatremia occurs frequently with secretion of antidiuretic hormone as a manifestation of the appropriate response of the hypothalamus and posterior pituitary to hypovolemia (Kaplowitz and Robertson, 1983a). Despite interstitial inflammation in the myocardium, left ventricular function is maintained, and in a small proportion of cases, the principal pathophysiologic effect on the heart is arrhythmia, presumably owing to vascular lesions involving the cardiac conduction system (Schmaier et al., 2001). Despite disseminated infection of the vascular endothelium, frequent occurrence of thrombocytopenia, and the stimulation of a procoagulant state, neither disseminated intravascular coagulation nor life-threatening hemorrhage occurs in rickettsial diseases except on rare occasions, as a remarkable manifestation of the homeostatic mechanisms of the coagulation system (Elghetany and Walker, 1999). The mechanisms of immune clearance of disseminated intraendothelial rickettsiae have been determined for SFG and TG rickettsiae (Valbuena et al., 2002). Rickettsial infection stimulates an early innate immune response with activation of natural killer cells and production of gamma interferon (IFN-γ), which act in concert to dampen rickettsial growth. Adaptive immunity develops with clonal expansion of CD4 and CD8 T lymphocytes and antibodyproducing B cells. Clearance of intraendothelial rickettsiae is achieved by rickettsicidal effects due to cytokine activation of the infected endothelial cells themselves. In mice, IFN-γ and tumor necrosis factor-α stimulate nitric oxide synthesis and nitric oxide-dependent rickettsial killing. In different human target cells activated by IFN-γ, tumor necrosis factor (TNF)-α, interleukin (IL)-Iβ, and RANTES (regulated on activation, normal T-cell expressed and secreted), rickettsicidal mechanisms include various combinations of nitric oxide, reactive oxygen species, and tryptophan degradation. Ultimate clearance of rickettsiae is mediated by major histocompatibility class (MHC) Imatched, cytotoxic CD8 T-lymphocyte activity that is largely perforin-dependent. The mechanisms by which the cytokine-secreting and cytotoxic cells are directed to the foci surrounding

CHAPTER 3.1.19

infected endothelium are important and undetermined. The production of particular chemokines in these foci suggests that the endothelium plays a key role. Remarkably, antibodies (including those directed at epitopes of OmpA and OmpB) also play a role in protective immunity. Rocky Mountain Spotted Fever Rocky Mountain spotted fever is the most severe rickettsiosis. In the pre-antibiotic era, 20–25% of previously healthy, infected persons died of the illness (Parker, 1941). Today, even with particular antimicrobial agents that are highly effective, 3–5% of persons die owing mainly to late or missed diagnosis and delayed or ineffective antimicrobial treatment (Dalton et al., 1995). The onset of disease follows an infective bite by a week (range 2–14 days), beginning with fever, severe headache, and myalgia. When patients present to a physician during the first three days of illness, only 3% have the classic triad of fever, rash, and history of tick bite (Helmick et al., 1984). The reasons are that up to 40% of patients are unaware of a tick bite, which is painless and may go unnoticed or be forgotten, and that the rash does not usually appear until day 3–5 of illness. To further confound the difficulty of diagnosis, symptoms such as nausea, vomiting, diarrhea, abdominal pain, and cough may suggest other diagnoses including enterocolitis, acute surgical abdomen, or pneumonia (Table 2). The rash typically appears on the ankles and wrists as faint pink 1–5 mm macules that represent a focus of vascular infection and surrounding vasodilation that blanches on pressure. These lesions may progress to become maculopapular, owing to the leakage of edema fluid from the affected blood vessels, with the development of a hemorrhage (petechia) in the center of the lesions in approximately half of cases. The rash will spread to involve the entire extremities, trunk and, in about half of the patients, the palms and soles. Skin necrosis or gangrene occurs as a result of rickettsial damage to the microcirculation in only 4% of patients and in some cases necessitates amputation (Kaplowitz et al., 1981; Kirkland et al., 1993; Hove and Walker, 1995). Just as the spread and growth of rickettsiae and damage to a greater portion of the vascular system occur in other organs, the involvement of the microcirculation of vital organs such as the lungs and brain causes life-threatening adult respiratory distress syndrome and encephalitis. In most fatal cases, death occurs 8–15 days after onset. If treatment is initiated during the first five days of symptoms, death is averted (Hattwick et al., 1978); however, in another clinical form, fulminant Rocky Mountain spotted fever, death

CHAPTER 3.1.19

The Order Rickettsiales

503

Table 2. Symptoms, signs, and laboratory data. Feature

RMSF

BF

% ATBF

ET

MT

Fever Headache Rash Tache noire Multiple eschars Myalgia Nausea and/or vomiting Abdominal pain Petechial rash Conjunctivitis Lymphadenopathy Stupor Diarrhea Edema Ataxia Meningismus Splenomegaly Hepatomegaly Jaundice Pneumonitis Cough Dyspnea Coma Seizures Shock and/or hypotension Decreased hearing Arrhythmia Myocarditis Death Increased AST level Thrombocytopenia Anemia Hyponatremia Azotemia

99–100 79–91 88–90 15 mm) and between 0 and 13.3% in small snails (8.0) that develops as the result of oxidation of the succinate. Bowdre et al. (1976) used a chemically defined liquid medium containing norepinephrine, which, like CHSS broth, supports growth of S. volutans under aerobic conditions. Growth of S. volutans on the surface of solid media (MPSS or CHSS broth solidified

CHAPTER 3.2.5

with 1.5% agar) is difficult but can be achieved by methods described by Padgett et al. (1982). The agar media must be supplemented with potassium metabisulfite (0.002%), catalase (230 units/ml), or SOD (30 units/ml). If catalase or SOD is used, it is added aseptically to the molten medium at 45°C just before dispensing into Petri dishes, whereas potassium metabisulfite is added prior to autoclaving. The medium is dispensed into Petri dishes (20 ml per dish) under dim illumination or red photographic safelight (red light does not cause photochemical generation of toxic forms of oxygen in the medium). The plates are allowed to dry in the dark at room temperature for 24 h. In dim illumination, 0.1-ml portions of appropriate dilutions of a broth culture of S. volutans are spread onto the agar surface with a glass rod. The plates are incubated for 5 days at 30°C in the dark in a sealed vessel containing 6– 12% O2 (the balance being N2). It is essential that the vessel be lined with moistened filter paper to maintain a high humidity. Even with these precautions, only 22–72% of the cells inoculated onto the plates develop into colonies. A method for obtaining reproducible colony counts was reported by Alban and Krieg (1996). The method uses a semisolid version of colony counting medium (CCM) as an overlay for a thicker layer of sterile medium. The medium contains pyruvate, which destroys hydrogen peroxide. Cells to be used for colony counts are grown in 5.0 ml of CCM broth contained in slanted 20 × 120-mm loosely screw-capped tubes incubated aerobically for 24 h at 30°C. Sterile semisolid medium (CCM plus 0.7% agar) is dispensed into Petri dishes (15 ml per dish) and allowed to gel for 30 min. A 0.1-ml volume of an appropriate dilution of the inoculum is added to 10 ml of semisolid CCM at 45°C and is poured onto the plates as an overlay. After this has gelled, the plates are incubated at 30°C in an atmosphere of 6% O2 and 94% N2 for 3–4 days.

Preservation Preservation of S. volutans can be accomplished by the following procedure (Pauley and Krieg, 1974): Cells from a turbid 24-hour-old broth culture are centrifuged at 3,500 × g and washed once in nutrient broth. The cells are then suspended by gentle agitation in a small amount of nutrient broth containing 10% dimethyl sulfoxide (DMSO). The resulting dense suspension is allowed to incubate for 30 min at room temperature and then 0.4-ml volumes are dispensed into plastic or glass vials suitable for liquid nitrogen preservation. The vials are sealed, frozen in a mixture of dry ice and alcohol, and then stored by submersion in liquid nitrogen. For recovery of

CHAPTER 3.2.5

the organisms, the vials are thawed in water at 30°C and transferred to semisolid MPSS or CHSS medium.

Physiology S. volutans has a strictly respiratory type of metabolism with oxygen as the terminal electron acceptor. It is microaerophilic in ordinary liquid media, such as nutrient broth (in which it grows poorly) and MPSS broth, and on solid media, such as CHSS agar supplemented with bisulfite. Also, it can grow aerobically in special liquid media such as CHSS broth or the chemically defined medium of Bowdre et al. (1976). Growth is inhibited by very low concentrations of H2O2 (0.29 µM or greater), and the organism is rapidly killed by concentrations of 10 µM or greater (Alban and Krieg, 1988). The microaerophilic nature of the organism has been attributed to the high susceptibility of the organism to toxic forms of oxygen which occur spontaneously in MPSS or CHSS broth under aerobic conditions, particularly when the media are illuminated (Padgett et al., 1982). Destruction of toxic forms of oxygen by media supplements such as SOD, catalase, potassium metabisulfite, or norepinephrine allows the organism to grow aerobically in broth (Padgett et al., 1982). However, even with these supplements, S. volutans can form colonies on MPSS, CHSS, or CCM agar only under microaerobic conditions (12% O2 or less). It will not grow even under microaerobic conditions unless the media supplements are present. Padgett and Krieg (1986) described two variant strains of S. volutans which, unlike the wild type, could grow in an air atmosphere (i.e., 21% O2) on CHSS agar containing potassium metabisulfite. Cell extracts of those aerotolerant variants exhibited greater o-dianisidine peroxidase activity than did the wild type, and this enzyme may be important for aerotolerance. The catalase-negative microaerophile Spirillum volutans is killed rapidly by levels of H2O2 greater than 10 µM. Alban and Krieg (1998) described a mutant isolated by single-step mutagenesis with diethyl sulfate that was able to survive and grow after exposure to 40 µM H2O2 and was effective in eliminating H2O2 added to the medium. Nevertheless, the mutant was no more tolerant to O2 than the wild type. The only apparent phenotypic difference between the wild type and the mutant was that the mutant had high NADH peroxidase activity (0.072 IU per mg protein) whereas the wild type had no detectable activity (90% of strains positive; −, >90% of strains negative; D, result varies among strains; and nd, no data available. From Etchebehere et al. (2001). b From Gumaelius et al. (2001). c As tested with auxanographic techniques using API systems. d From Tamaoka et al. (1987). e From Willems et al. (1989). a

can be added here that the presence of hydroxy putrescine is characteristic for the genera of the β-Proteobacteria. Table 3 gives features for the differentiation of Comamonas species.

Cultivation Comamonas strains can be grown on nutrient agar and many other commonly used media such as tryptic soy agar and Columbia agar with or without blood. They are able to use many organic acids and amino acids, but few sugars (De Vos et al., 1985). They can degrade a variety of complex aromatic compounds, steroids and many manmade complex organic compounds. Optimal growth temperature is 30°C.

Preservation Comamonas cultures can be maintained on nutrient agar slants at 4°C for up to 2 months. Lyophilization or storing in liquid nitrogen can be used for long-term preservation of strains on the condition that cryoprotectants are added. Generally, the use of standard procedures is recommended.

Physiology Comamonas strains have an aerobic respiratory type of metabolism. Most strains of C. terrigena and C. testosteroni are capable of nitrate reduc-

tion, but cannot reduce nitrites (Willems et al., 1991), while all strains of C. nitrativorans and C. denitrificans can reduce nitrate all the way to N2. Although Comamonas strains were identified in a study of denitrifying biofilms at a water treatment plant, they were thought not to be true members of the biofilm community, but to have washed in from previous stages (Lemmer et al., 1997). Other data, however, confirm the denitrification capacity of some members of the genus (Vanbrabant et al., 1993). In particular, it has been shown that Comamonas strain SGLY2 is capable of aerobic denitrification and nitrogen production without nitrite building up. This strain has been studied for use in mixed-culture aerated reactors (Patureau et al., 1997). Comamonas strains are able to degrade a wide variety of aromatic compounds. Comamonas strain JS765 has a 1,2-dioxygenase able to break down nitrobenzene to nitrohydrodiol which spontaneously decomposes to catechol and nitrite. Catechol is then broken down via the meta-cleavage pathway, the genes of which have been sequenced (Parales et al., 1997; Genetics). This pathway has been studied in comparison with similar pathways in Pseudomonas strains (He and Spain, 1999). Comamonas testosteroni strains that degrade polycyclic aromatic compounds such as phenanthrene, naphthalene and anthracene have been described and the genes involved Genetics have been characterized (Goyal and Zylstra, 1996). The mechanism for phenol tolerance was studied in the phenoldegrading C. tesosteroni strains P15 and E23 (Yap et al., 1999).

CHAPTER 3.2.7

The meta-cleavage pathway is used by C. testosteroni for the degradation of 4-chlorophenol and related compounds (Hollender et al., 1997). Comamonas testosteroni can degrade 4-toluene sulfonate, which is thought to be taken up by an inducible secondary proton symport system (Locher et al., 1993). The degradation of 4toluene sulfonate and 4-toluene carboxylate involves oxygenation of the side chain to 4sulfobenzoate and terephthalate, respectively, which are then oxidized to protocatechuate. Genes involved in these pathways have been sequenced and characterized (Junker et al., 1997b; Genetics). Terephthalate is broken down by a terephthalate dioxygenase system to (1R, 2S)-dihydroxy-3,5-cyclohexadiene-1,4-dicarboxylic acid which is further degraded to protocatechuate by (1R,2S)-dihydroxy-1,4dicarboxy-3,5-cyclohexadiene dehydrogenase (Oppenberg et al., 1995). The dioxygenase system is a Rieske [2Fe-2S] protein with two subunits α and β which are thought to form an α2β2 structure (Schlaefi et al., 1994). The sulfonate and sulfate ester degradation as a source of sulfur for growth of Gram-negative bacteria has recently been reviewed (Kertesz, 2000). Comamonas testosteroni I2 was shown to degrade recalcitrant components such as aniline and chloroaniline (Boon et al., 2001). Both components are among the most widely produced industrial amines commonly used for the production of polyurethanes, rubber, azo dyes, drugs, pesticides, etc. The presence of these toxic amines in natural environments is strictly regulated by the environmental protection agencies of the European Union (EU) and the United States. Testosterone can be used as sole carbon source by C. testosteroni and induces the expression of enzymes involved in catabolism of steroids and aromatic hydrocarbons and the repression of at least one amino acid degrading enzyme. It was suggested that steroids may play a regulatory role in the synthesis of catabolic enzymes during adaptive growth (Moebus et al., 1997). Several of the enzymes involved in the conversion and break down of steroids have been studied and their genes sequenced (Genetics). The structure and function of δ5-3-ketosteroid isomerase and 3-oxo-δ5-steroid isomerase, which catalyze the the conversion of δ5- to δ4-3-ketosteroids by intramolecular proton transfer, have been studied (Brothers et al., 1995; Zao et al., 1996, 1997). A bile acid 3-α-sulfate sulfohydrolase of C. testosteroni was purified and characterized (Tazuke et al., 1994). Comamonas strains enriched from wastewater from an aerated stabilization basin of a bleached kraft pulp mill were reported to degrade terpenes such as abietane and pimarane-type resin acids (Morgan and Wyndham, 1996).

Comamonas

729

Some Comamonas strains are able to degrade poly-3-hydroxybutyrate using an extracellular poly-3-hydroxybutyrate depolymerase, which can also hydrolyze poly(3-hydroxybutyrate-co3-hydroxyvalerate) and poly-3-hydroxyvalerate. The enzyme differs from other poly-3-hydroxybutyrate depolymerases in that it is insensitive to phenylmethylsulfonyl fluoride and hydrolyzes poly-3-hydroxybutyrate to 3-hydroxybutyrate monomers (Jendrossek et al., 1993). The encoding gene of a Comamonas sp. strain was cloned and partly sequenced, and the resulting partial protein was analyzed (Jendrossek et al., 1995). A very similar sequence was obtained for C. testosteroni strain YM1004 (Shinomiya et al., 1997). Comamonas sp. strain P37C, isolated from compost, was able to degrade a range of poly-3hydroxyalkanoates (Quinteros et al., 1999). The enzymes involved in the first steps of quinoline and 3-methyl quinoline degradation by C. testosteroni strain 63, quinoline 2-oxidoreductase and 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase, have been characterized. Quinoline 2oxidoreductase is a molybdo-iron flavoprotein and 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase is a single component protein (Schach et al., 1995). Comamonas testosteroni produces 5-aminolevulinic acid, a precursor to tetrapyrroles, from aminoacylated tRNA-Glu via a two-step pathway involving glutamyl-tRNA reductase and glutamine-1-semialdehyde-2,1-aminomutase (Hungerer et al., 1995). The biosynthesis of aromatic amino acids via several pathways has been extensively studied among pseudomonad bacteria. In C. testosteroni prephenate dehydrogenase and arogenate dehydrogenase are reactive with either nicotinamide adenine dinucleotide (NAD) or nicotinamide adenine dinucleotide phosphate (NADP). No arogenate dehydratase activity is present; prephenate dehydratase is present and its activity is stimulated by L-tyrosine (Byng et al., 1983).

Genetics The genes of C. testosteroni involved in the degradation of naphthalene and phenanthrene have been cloned and characterized. At least two different sets of phenanthrene degradation genes may be present among different Comamonas testosteroni strains. They are different from the naphthalene degradation genes (nha) of Pseudomonas putida (Goyal and Zylstra, 1996). The gene phtD for 4,5-dihydroxyphthalate decarboxylase, an enzyme involved in phthalate metabolism in C. testosteroni, has been identified and sequenced. The deduced amino acid sequence shows similarity to the pth5 genes of P.

730

A. Willems and P. De Vos

putida, and the sequence upstream of pthD shows a high similarity to pth1 in P. putida, where it is thought to be a positive regulator of the other pth genes. These findings suggest a common origin for the genes involved in the phthalate metabolism (Lee et al., 1994). The bphD gene of C. testosteroni strain B-356, encoding for 2-hydroxy-6-oxo (phenyl/chlorophenyl) hexa-2,4-dienoic acid hydrolase, was sequenced, and the gene product has a mechanism of action similar to that of classical lipases and hydrolases (Ahmad et al., 1995). In the same strain, biphenyl/chlorobiphenyl dioxygenase, the enzyme system involved in the first step of the degradation of biphenyl or chlorobiphenyl compounds, was characterized. The genes for the three components of this enzyme system have been sequenced: bphA and bphE encode the two subunits of a terminal Fe-S oxygenase, bphF encodes a ferredoxin, and bphG encodes a ferredoxin reductase and is not located near the bphAEF genes (Sylvestre et al., 1996b). Many of these enzymes have been characterized (Hurtubise et al., 1995; Hurtubise et al., 1996) and some have been used to produce enzyme chimeras by combination with oxygenase components from other bacteria (Chebrou et al., 1999). Genes for subsequent degradation steps have also been characterized: bphB encodes 2,3dihydro-2,3-dihydroxybiphenyl dehydrogenase (Sylvestre et al., 1996a) and bphC encodes 2,3dihydroxybiphenyl-1,2-dioxygenase which catalyzes meta-1,2 fission of the aromatic ring (Bergeron et al., 1994). Finally, it has been demonstrated (Hein et al., 1998) that in this strain two meta-cleavage dioxygenases coexist with different degrees of sensitivity towards intermediates of the biphenyl catabolic pathway and suggesting a complex regulation system. Genes involved in the degradation of catechol via the meta-cleavage pathway have been sequenced and amino acid sequences deduced for strain Comamonas sp. JS765: cdoE encodes catechol 2,3-dioxygenase, cdoT encodes a ferredoxin, and cdoR is a regulatory gene (Parales et al., 1997). A gene cluster encoding for the degradation of 3-(3-hydroxyphenyl)-propionate from C. testosteroni strain TA441 was cloned and sequenced. It contains a regulatory gene, mhpR, and the following six genes for catabolic enzymes: mhpA for a flavin-type hydroxylase, mhpB for an extradiol dioxygenase, mhpD for 2-keto-4-pentenoate hydratase, mhpF for an acylating acetaldehyde dehydrogenase, mhpE for 4-hydroxy-2-ketovalerate aldolase, and mhpC for a meta-cleavage hydrolase (Arai et al., 1999b). The same research group (Arai et al., 2000) studied the arrangement and the regulation of the meta pathway in the breakdown of phenol.

CHAPTER 3.2.7

The genes involved in the degradation of 4toluene sulfonate and 4-toluene carboxylate of C. testosteroni have been extensively studied. The first step of oxygenation of the side chain to 4-sulfobenzoate and terephthalate, respectively, requires three enzymes: a 4-toluenesulfonate methyl-monooxygenase system consisting of a reductase B and oxygenase M encoded by the tsaBM genes, a 4-sulfobenzyl alcohol dehydrogenase encoded by tsaC and a 4-sulfobenzaldehyde encoded by tsaD. These genes are coexpressed under the regulation of the product of tsaR. Some of the enzymes involved have been purified and partially characterized. The tsaMB oxygenase system is a class IA mononuclear iron oxidase with tsaM having a Rieske [2Fe-2S] center, and tsaC is a short-chain zinc-dependent dehydrogenase (Junker et al., 1996). The conversion of 4-sulfobenzoate to protocatechuate is catalyzed by a 4-sulfobenzate-3,4-dioxygenase system consisting of a reductase C and an oxygenase A, again with a Rieske [2Fe-2S] center, encoded by the genes psbAC (Junker et al., 1996). The genes tsaMBCD, tsaR, and psbAC are located on conjugative plasmids in some C. testosteroni strains (Junker and Cook, 1997a). Terephthalate is also broken down to protocatechuate, but genes involved have not yet been characterized. Protocatechuate is broken down via a meta-cleavage pathway by enzymes thought to be encoded on the chromosome (Junker and Cook, 1997a). The gene cluster encoding a phenol hydroxylase in the phenol-degrading C. testosteroni strain R5 (phcKLMNOP) and the regulatory gene phcR have been cloned and expressed in Pseudomonas aeruginosa, conferring the use of phenol as sole carbon source on this bacterium (Teramoto et al., 1999). A second gene cluster which can confer the ability to use phenol (aphKLMNOPQB and the corresponding regulatory genes aphR and aphS) was studied in C. testosteroni strain TA441 (Arai et al., 1999a). In C. testosteroni, the genes for several enzymes involved in the breakdown of steroid compounds have been studied. The gene for a 3-α-hydroxysteroid dehydrogenase has been sequenced and expressed through cloning, and the enzyme was characterized (Abalain et al., 1995; Oppermann and Maser, 1996a). Because of its similarity to ribosomal proteins L10 and L7/ 12, it was suggested that this enzyme may be formed by fusion of two ribosomal proteins (Baker, 1996). The gene for 3-ketosteroid-δ4-5-αdehydrogenase was sequenced and located downstream of a gene for 3-ketosteroid-δ1dehydrogenase. Although both enzymes are functionally similar, their genes are probably not derived from a common ancestor (Florin et al., 1996). A 3β/17β hydroxysteroid dehydrogenase

CHAPTER 3.2.7

was studied by mutagenic replacement within the active site to identify important residues (Oppermann et al., 1997). The steroid-inducible gene stdC of C. testosteroni was cloned and sequenced. It was homologous to open reading frames (ORFs) of unknown function in the polyhydroxyalkanoic acid cluster of several other bacteria and is transcribed as a monocistronic mRNA that is abundant in cells grown on steroid carbon sources (Cabrera et al., 1997). Recently, in a model for the regulation of the 3-αhydroxysteroid hydrogenase/carbonyl reductase (Xiong et al., 2001), it was proposed that the hydrogenase is under the control of two repressor proteins of which the binding to the promotor and the messenger RNA is de-repressed by an inducer such as testosterone. The gene for an aliphatic nitrilase from C. testosteroni was sequenced and overexpressed, and the enzyme was purified and its primary structure determined. A Cys163 residue is thought be essential in the active site (Levy-Schil et al., 1995). Several plasmids have been isolated from or transferred to Comamonas strains, and they may contribute to the catabolic versatility of these organisms. Comamonas testosteroni strains T-2 and PSB-4 and the type strain were reported to contain plasmids pTSA and pT2T, plasmid pPSB, and no plasmids, respectively (Junker and Cook, 1997a). Plasmid pTSA (85 kb) is a conjugative plasmid of the IncP1 group and carries the genes tsaMBCD, tsaR and psbAC, involved in the degradation of 4-toluene sulfonate to protocatechuate, and two copies of insertion sequence IS1071. Plasmid pPSB (85 kb) also is a conjugative plasmid; it carries only the psbAC gene as well as two copies of IS1071. Conjugative experiments with the type strain have suggested that the psb genes are located in a composite transposon (Junker and Cook, 1997a). Plasmid RP4::Tn4371 with genes for biphenyl and 4-chlorobiphenyl degradation was transferred from Enterobacter agglomerans to indigenous soil bacteria, including Comamonas strains, where these catabolic enzymes were successfully expressed (De Rore et al., 1994). Plasmid PR4::Mu3A, carrying chromosomal DNA fragments encoding for the use of biphenyl as a sole carbon source, were transferred to C. testosteroni and the enzymes were effectively expressed (Springael et al., 1996). Plasmids pACK5 and pACT72, including a replicon of the cryptic Acetobacter pasteurianus plasmid pAC1, were transferred to and successfully expressed in C. terrigena (Grones and Turna, 1995). Plasmid pE43 from Pseudomonas aeruginosa and plasmid pPC3 from Arthrobacter globiformis, carrying genes for dehalogenation of chlorobenzoates, permitted C. testosteroni strain VP44 to grow on

Comamonas

731

ortho- or para-biphenyls as sole carbon source (Hrywna et al., 1999). Plasmid PNB2 (Boon et al., 2001) has been isolated from Comamonas testosteroni I2, an aniline- and chloroaniline-degrading strain. So far, the chloroaniline capacity could not be demonstrated on the plasmid by conjugative transformation with a Ralstonia strain, in contrast to the aniline capacity. Additional data suggest that oxidative deamination of chlorinated and nonchlorinated aromatics may be completed by a different set of genes (Boon et al., 2001).

Ecology Comamonas strains have been recovered from sites heavily contaminated with complex organic compounds and heavy metals, and resistance to heavy metals such as cadmium and nickel has been reported (Stoppel and Schlegel, 1995; Kanazawa and Mori, 1996). A strain tentatively identified as C. testosteroni was among the strongest chromate-reducing bacteria found in the cooling water of an electricity plant and may therefore be implicated in the blockage of pipes through precipitation of chromium (III) oxide (Cooke et al., 1995). Most of the isolates from a polychlorinated biphenyl (PCB)-polluted site were Comamonas testosteroni strains capable of degrading biphenyl and a variety of polychlorinated biphenyls (Joshi and Walia, 1995). Many Comamonas strains from various environments are capable of degrading poly-3hydroxybutyrate (PHB; Mergaert and Swings, 1996), and the extracellular PHB depolymerase of one strain has been purified and characterized (Jendrossek et al., 1993). The broad substrate specificity of its 3-αhydroxysteroid dehydrogenases and its occurrence in the intestinal tract of vertebrates have led to suggestions that C. testosteroni and related bacterial strains may contribute to the bioactivation or inactivation of hormones, bile acids and xenobiotics (Oppermann and Maser, 1996a). Comamonas testosteroni possesses steroidinducible hydroxysteroid dehydrogenases and carbonyl reductases which may provide protection against natural and synthetic toxic compounds in soil and the intestinal tract of mammals (Oppermann et al., 1996b).

Pathogenicity Although many Comamonas strains have been isolated from clinical samples and from the hospital environment (Willems et al., 1991), they are regarded as very occasional opportunistic pathogens (Gilardi, 1985), and no evidence of

732

A. Willems and P. De Vos

pathogenic effect on healthy people has been reported.

Applications Because they can break down a wide variety of complex organic compounds (including carboxylic and dicarboxylic aliphatic or unsaturated carboxylic acids, aromatic compounds, and sterols), Comamonas strains contribute to bioremediation processes. Comamonas testosteroni has been used for the degradation of 4-toluenesulfonic acid in a continuously operated fixed-bed biofilm reactor (Khlebnikov and Peringer, 1996). Analysis of the bacterial communities in activated sludge revealed the presence of Comamonas strains (Kaempfer et al., 1996). Comamonas terrigena strain N3H, immobilized in an alginate gel, has been used for the degradation of the anion-active surfactant dihexyl-sulfosuccinate (Huska et al., 1996; Toth et al., 1996; Proksova et al., 1999). Comamonas strains JS46 and JS 47 have been studied in mixed and non-mixed culture immobilized cell reactors for the degradation of m- and p-nitrobenzoate (Goodall et al., 1998b). The experimental data were used for process modeling (Goodall and Peretti, 1998a). A quinohemoprotein ethanol dehydrogenase, which catalyzes the NAD-independent oxidation of a broad range of alcohols to their aldehydes and on to the carboxylic acid, has been purified from ethanol-grown C. testosteroni cells. The enzyme was studied for use in the production of (S)-solketal (2,2-dimethyl-1,3-dioxolane-4methanol) by enantioselective oxidation of racemic solketal (Geerlof et al., 1994; Machado et al., 1999). The active holoenzyme contains pyrroquinoline-quinone, calcium ions, and heme c (De Jong et al., 1995). The encoding gene (qhedh) was cloned, sequenced and expressed in E. coli (Stoorvogel et al., 1996). The enzyme has been applied in electrodes through immobilization in a redox polymer network (Stigter et al., 1997) and can be used for the enantioselective oxidation of secondary alcohols (Godde et al., 1999). The capacity of C. testosteroni BS1310 (pBS1010) to degrade sulfoaromatic compounds resulted in the design of a Clark-type electrode to measure the p-toluene concentration (Makarenko et al., 1999). The ability of C. testosteroni to selectively degrade L-lysine from racemized lysine crystals has been applied in the large-scale production of D-lysine (Takahashi et al., 1997). Finally, a number of studies point to other potential applications of Comamonas strains. Comamonas testosteroni strain CMI 2848 had the highest activity for hydroxylating 3cyanopyridine to 3-cyano-6-hydroxypyridine

CHAPTER 3.2.7

among 4600 isolates screened in a study contributing to the synthesis of new agricultural chemicals (Yasuda et al., 1995). Because hydroxylated N-hetero-aromatic compounds are difficult to produce, the characteristic of a quinolineoxidoreductase of C. testosteroni 63 to mediate the hydroxylation of N-hetero-aromatic compounds such as pyridine to hydroxy-substituted pyrimidines may be of pharmaceutical importance (Fetzner, 1999).

Literature Cited Abalain, J. H., S. Di Stefano, M. L. Abalain-Colloc, and H. H. Floch. 1995. Cloning, sequencing and expression of Pseudomonas testosteroni gene encoding 3alpha-hydroxysteroid dehydrogenase. J. Steroid Biochem. Molec. Biol. 55:233–238. Ahmad, D., J. Fraser, M. Sylvestre, A. Larose, A. Kahn, J. Bergeron, J. M. Juteau, and M. Sondossi. 1995. Sequence of the bphD gene encoding 2hydroxy-6-oxo-(phenyl/ chlorophenyl)hexa-2,4 dienoic acid (HOP/cPDA) hydrolase involved in the biphenyl/polychlorinated biphenyl degradation pathway in Comamonas testosteroni: Evidence suggesting involvement of Ser112 in catalytic activity. Gene 156:69–74. Amann, R., W. Ludwig, R. Schulze, S. Spring, E. Moore, and K.-H. Schleifer. 1996. rRNA-targeted oligonucleotide probes for the identification of genuine and former pseudomonads. Syst. Appl. Microbiol. 19:501–509. Arai, H., Akahira, S., T. Ohishi, and T. Kudo. 1999a. Adaptation of Comamonas testosteroni TA441 to utilization of phenol by spontaneous mutation of the gene for a trans-acting factor. Molec. Microbiol. 33:1132–1140. Arai, H., T. Yamamoto, T. Ohishi, T. Shimizu, T. Nakata, and T. Kudo. 1999b. Genetic organization and characteristics of the 3-(3-hydroxyphenyl)propionic acid degradation pathway of Comamonas testosteroni TA441. Microbiology 145:2813–2820. Arai, H., T. Ohishi, M. Y Chang, and T. Kudo. 2000. Arrangment and regulation of the genes for the meta-pathway enzymes required for the degradation of phenol in Comamonas testosteroni TA441. Microbiology 146: 1707–1715. Baker, M. E. 1996. Bacterial 3-alpha-hydroxysteroid dehydrogenase is homologous to a fusion of bacterial ribosomal L10 and L7/12 genes. J. Steroid Biochem. Molec. Biol. 59:365–366. Ben-Tovim, T., E. Eylan, A. Romano, and R. Stein. 1974. Gram-negative bacteria isolated from external eye infections. Infection 2:162–165. Bergeron, J., D. Ahmad, D. Barriault, A. Larose, M. Sylvestre, and J. Powlowski. 1994. Identification and mapping of the gene translation products involved in the first steps of the Comamonas testosteroni B-356 biphenyl/ chlorobiphenyl degradation pathway. Can. J. Microbiol. 40:743–753. Boon, N., J. Goris, P. De Vos, W. Verstraete, and E. Top. 2000. Bioaugmentation of activated sludge by an indigenous 3-chloroaniline-degrading Comamonas testosteroni strain, I2gfp. Appl. Environ. Microbiol. 66:2906– 2913. Boon, N., J. Goris, P. De Vos, W. Verstraete, and E. Top. 2001. Genetic diversity among 3-chloroaniline- and aniline

CHAPTER 3.2.7 degrading strains of the Comamonadaceae. Appl. Environ. Microbiol. 67:1107–1115. Brothers, P. N., G. Blotny, L. Qi, and R. M. Pollack. 1995. An active site phenylalanine of 3-oxo-d5-steroid isomerase is catalytically important for proton transfer. Biochemistry 34:15453–15458. Byng, G. S., J. L. Johnson, R. J. Whitaker, R. L. Gherna, and R. A. Jensen. 1983. The evolutionary pattern of aromatic amino acid biosynthesis and the emerging phylogeny of pseudomonad bacteria. J. Molec. Evol. 19:272–282. Cabrera, J. E., G. Panzetta-Dutari, J. L. Pruneda, and S. Genti-Raimondi. 1997. A new Comamonas testosteroni steroid-inducible gene: cloning and sequence analysis. J. Steroid Biochem. Molec. Biol. 63:91–98. Chebrou, H., Y. Hurtubise, D. Barriault, and M. Sylvestre. 1999. Heterologous expression and characterization of the purified oxygenase component of Rhodococcus globerulus P6 biphenyl dioxygenase and of chimeras derived from it. J. Bacteriol. 181:4805–4811. Cooke, V. M., M. N. Hughes, and R. K. Poole. 1995. Reduction of chromate by bacteria isolated from the cooling water of an electricity generating station. J. Indust. Microbiol. 14:323–328. Coote, J. G., and H. Hassall. 1973. The degradation of Lhistidine, imidazolyl-L-lactate, and imidazolylpropionate by Pseudomonas testosteroni. Biochem. J. 132:409–422. Dagley, S., and M. D. Patel. 1957. Oxidation of p-cresol and related compounds by a Pseudomonas. Biochem. J. 66:227–233. Davis, G. H. G., and R. W. A. Park. 1962. A taxonomic study of certain bacteria currently classified as Vibrio species. J. Gen. Microbiol. 27:101–119. De Jong, G. A. H., A. Geerlof, J. Stoorvogel, J. A. Jongejan, S. De Vries, and J. A. Duine. 1995. Quinohaemoprotein ethanol dehydrogenase from Comamonas testosteroni: Purification, characterization and reconstitution of the apoenzyme with pyrroloquinoline quinone analogues. Eur. J. Biochem. 230:899–905. De Rore, H., K. Demolder, K. De Wilde, E. Top, F. Houwen, and W. Verstraete. 1994. Transfer of the catabolic plasmid RP4::Tn4371 to indigenous soil bacteria and its effect on respiration and biphenyl breakdown. FEMS Microbiol. Ecol. 15:71–77. De Vos, P., K. Kersters, E. Falsen, B. Pot, M. Gillis, P. Segers, and J. De Ley. 1985. Comamonas Davis and Park 1962 gen. nov., nom. Rev. emend., and Comamonas terrigena Hugh 1962 sp. nov., nom. rev. Int. J. Syst. Bacteriol. 35:443–453. Etchebehere, C., M. I. Errazquin, P. Dabert, R. Moletta, and L. Muxí. 2001. Comamonas nitrativorans sp. nov., a novel denitrifier isolated from a denitrifying reactor treating landfill leachate. Int. J. Syst. Evol. Microbiol. 51:977–983. Fetzner, S. 1999. Bioconversion of pyrimidine by resting cells of quinoline-degrading bacteria. FEMS Microbiol. Lett. 176:291–299. Florin, C., T. Koehler, M. Grandguillot, and P. Plesiat. 1996. Comamonas testosteroni 3-ketosteroid-δ4(5-α)dehydrogenase gene and protein characterization. J. Bacteriol. 178:3322–3330. Galarneault, T. P., and E. Leifson. 1956. Taxonomy of Lophomonas N. Gen. Can. J. Microbiol. 2:102–110. García-Valdés, E., E. Cozar, R. Rotger, J. Lalucat, and J. Ursing. 1988. New naphthalene-degrading marine Pseudomonas strains. Appl. Environ. Microbiol. 54:2478–2485.

Comamonas

733

Geerlof, A., J. Stoorvogel, J. A. Jongejan, E. J. T. M. Leenen, T. J. G. M. Van Dooren, W. J. J. Van Den Tweel, and J. A. Duine. 1994. Studies on the production of (S)-(+)solketal (2,2-dimethyl-1,3-dioxolane-4-methanol) by enantioselective oxidation of racemic solketal with Comamonas testosteroni. Appl. Microbiol. Biotechnol. 42:8–15. Gilardi, G. L. 1971. Characterization of nonfermentative, nonfastidious Gram-negative bacteria encountered in medical bacteriology. J. Appl. Bacteriol. 34:623–644. Gilardi, G. L. 1985. Pseudomonas. In: E. H. Lennette, A. Balows, W. J. Hausler Jr., and H. J. Shadomy (Eds.) Manual of Clinical Microbiology, 4th ed. American Society for Microbiology. Washington, DC. 350–372. Godde, C., M. Liebergesell, and A. Steinbuchel. 1999. Isolation of poly(beta-L-malic acid)-degrading bacteria and purification and characterization of the PNA hydrolase from Comamonas acidovorans strain 7789. FEMS Microbiol. Lett. 173:365–372. Goodall, J. L., and S. W. Peretti. 1998a. Dynamic modeling of meta- and para-nitrobenzoate metabolism by a mixed co-immobilized culture of Comamonas spp. JS46 and JS47. Biotech. Bioeng. 59:507–516. Goodall, J. L., S. M. Thomas, J. C. Spain, and S. W. Peretti. 1998b. Operation of mixed-culture immobilized cell reactors for the metabolism of meta- and paranitrobenzoate by Comamonas sp. JS46 and Comamonas sp. JS47. Biotechnol. Bioeng. 59:21–27. Goyal, A. K., and G. J. Zylstra. 1996. Molecular cloning of novel genes for polycyclic aromatic hydrocarbon degradation from Comamonas testosteroni GZ29. Appl. Environ. Microbiol. 62:230–236. Gray, P. H. H., and H. G. Thornton. 1928. Soil bacteria that decompose certain aromatic compounds. Zentralbl. Bakteriol. Parasitenk. Abt. 2 73:74–96. Grones, J., and K. Turna. 1995. Transformation of microorganisms with the plasmid vector with the replicon from pAC11 from Acetobacter pasteurianus. Biochem. Biophys. Res. Comm. 206:942–947. Gumaelius, L., G. Magnusson, B. Petterson, and G. Dalhammar. 2001. Comamonas denitrificans sp.nov., an efficient denitrifying bacterium isolated from activated sludge. Int. J. Syst. Evol. Microbiol. 51:999–1006. Gunther, K. A. 1894. Über einen neuen in Erdboden gefundenen Kommabacillus. Zentralbl. Bakteriol. Parasitenkd. 16:746–747. He, Z. Q., and J. C. Spain. 1999. Comparison of the downstream pathways for degradation of nitrobenzene by Pseudomonas pseudoalcaligenes JS45 (2-aminophenol pathway) and by Comamonas sp. JS765 (catechol pathway). Arch. Microbiol. 171:309–316. Hein, P., J. Powlowski, D. Barriault, Y. Hurtubise, D. Ahamad, and M. Sylvestre. 1998. Biphenyl-associated meta-cleavage dioxygenase from Comamonas testosteroni B-356. Can. J. Microbiol. 44:42–49. Hollender, J., W. Hopp, and W. Dott. 1997. Degradation of 4-chlorophenol via the meta cleavage pathway by Comamonas testosteroni JH5. Appl. Environ. Microbiol. 63:4567–4572. Hrywna, Y., T. V. Tsoi, O. V. Maltseva, J. F. Quensen, and J. M. Tiedje. 1999. Construction and characterization of two recombinant bacteria that grow on ortho- and parasubstituted biphenyls. Appl. Environ. Microbiol. 65:2163–2169.

734

A. Willems and P. De Vos

Hugh, R. 1962. Comamonas terrigena comb. nov. with proposal of a neotype an request for an opinion. Int. Bull. Bacteriol. Nomencl. Tax. 12:33–35. Hugh, R. 1965. A comparison of Pseudomonas testosteroni and Comamonas terrigena. Int. Bull. Bacteriol. Nomencl. Tax. 15:125–132. Hungerer, C., B. Troup, U. Roemling, and D. Jahn. 1995. Regulation of the hemA gene during 5-aminolevulinic acid formation in Pseudomonas aeruginosa. J. Bacteriol. 177:1435–1443. Hurtubise, Y., D. Barriault, J. Powlowski, and M. Sylvestre. 1995. Purification and characterization of the Comamonas testosteroni B-356 biphenyl dioxygenase components. J. Bacteriol. 177:6610–6618. Hurtubise, Y., D. Barriault, and M. Sylvestre. 1996. Characterization of active recombinant His-tagged oxygenase component of Comamonas testosteroni B-356 biphenyl dioxygenase. J. Biol. Chem. 271:8152–8156. Huska, J., I. Zavdska, D. Toth, M. Dobrotova, and P. Gemeiner. 1996. Immobilization of surfactant degrading bacteria in alginate gel. Biologia (Bratislava) 51:279– 283. Jayarao, B. M., and L. Wang. 1999. A study on the prevalence of gram-negative bateria in bulk tank milk. J. Dairy Sci. 82:2620–2624. Jendrossek, D., I. Knoke, R. B. Habibian, A. Steinbüchel, and H. G. Schlegel. 1993. Degradation of poly(3-hydroxybutyrate), PHB, by bacteria and purification of a novel PHB depolymerase from Comamonas sp. J. Environ. Polymer Degrad. 1:53–63. Jendrossek, D., M. Backhaus, and M. Andermann. 1995. Characterization of the extracellular poly(3-hydroxybutyrate) depolymerase of Comamonas sp. and of its structural gene. Can. J. Microbiol. 41:160–169. Joshi, B., and S. Walia. 1995. Characterization by arbitrary primer polymerase chain reaction of polychlorinated biphenyl (PCB)-degrading strains of Comamonas testosteroni isolated from PCB-contaminated soil. Can. J. Microbiol. 41:612–619. Junker, F., E. Saller, H. R. S. Oppenberg, P. M. H. Kroneck, T. Leisinger, and A. M. Cook. 1996. Degradative pathways for p-toluenecarboxylate and p-toluenesulfonate and their multicomponent oxygenases in Comamonas testosteroni strains PSB-4 and T-2. Microbiology 142:2419–2427. Junker, F., and A. M. Cook. 1997a. Conjugative plasmids and the degradation of arylsulfonates in Comamonas testosteroni. Appl. Environ. Microbiol. 63:2403–2410. Junker, F., R. Kiewitz, and A. M. Cook. 1997b. Characterization of the p-toluenesulfonate operon tsaMBCD and tsaR in Comamonas testosteroni T-2. J. Bacteriol. 179:919–927. Kaempfer, P., R. Erhart, C. Beimfohr, J. Boehringer, M. Wagner, and R. Amann. 1996. Characterization of bacterial communities from activated sludge: Culture-dependent numerical identification versus in situ identification using group- and genus-specific rRNA-targeted oligonucleotide probes. Microb. Ecol. 32:111–121. Kanazawa, S., and M. Mori. 1996. Isolation of cadmiumresistant bacteria and their resistance mechanisms. Part I: Isolation of CD-resistant bacteria from soils contaminated with heavy metals. Soil Sci. Plant Nutr. 42:725– 730. Kertesz, M. A. 2000. Riding the sulfur cycle—metabolism of sulfonates and sulfate esters in Gram-negative bacteria. FEMS Microbiol. Rev. 24:135–175.

CHAPTER 3.2.7 Khlebnikov, A., and P. Peringer. 1996. Biodegradation of ptoluenesulphonic acid by Comamonas testosteroni in an aerobic counter-current structured packing biofilm reactor. Water Sci. Technol. 34:257–266. Krieg, N. R., and J. G. Holt (Eds.). 1984. Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. Laitinen, S., M. Linnainmaa, J. Laitinen, H. Kiviranta, M. Reiman, and J. Liesivuori. 1999. Endotoxins and IgG antibodies as indicators of occupational exposure to the microbial contaminants of metal-working fluids. Int. Arch. Occup. Environ. Health. 72:443–450. Lee, J.-H., T. Omori, and T. Kodama. 1994. Identification of the metabolic intermediates of phthalate by Tn5 mutants of Pseudomonas testosteroni and analysis of the 4,5-dihydroxyphthalate decarboxlase gene. J. Ferment. Bioeng. 77:583–590. Lehmann, K. B., and R. Neumann. 1927. Bakteriologie insbesondere bakteriologische Diagnostiek. II. Allgemeine und spezielle Bakteriologie, 7th ed. J. F. Lehmann. Munich, Germany. 548. Lemmer, H., A. Zaglauer, A. Neef, H. Meier, and R. Amann. 1997. Denitrification in a methanol-fed fixedbed reactor. Part 2: Composition and ecology of the bacterial community in the biofilms. Water Res. 31:1903–1908. Levy-Schil, S., F. Soubrier, A.-M. Crutz-Le Coq, D. Faucher, J. Crouzet, and D. Petre. 1995. Aliphatic nitrilase from a soil isolated Comamonas testosteroni sp.: Gene cloning and overexpression, purification and primary structure. Gene 161:15–20. Locher, H. H., Poolman, B., Cook, A. M., and Konings, W. N. 1993. Uptake of 4-toluene sulfonate by Comamonas testosteroni T-2. J. Bacteriol. 175:1075–1080. Machado, S. S., A. Jongejan, A. Geerlof, J. A. Jongejan, and J. A. Duine. 1999. Entropic and enthalpic contributions to the enantioselectivity of quinohaemoprotein alcohol dehydrogenases from Acetobacter pasteurianus and Comamonas testosteroni in the oxidation of primary and secondary alcohols. Biocatal. Biotrans. 17:179–207. Makarenko, A. A., S. V. Balashov, T. N. Kuvichkina, P. V. Il’yasov, and A. N. Reshetilov. 1999. A biosensor of sulfoaromatic compounds based on the strain Comamonas testosteroni BS1310 (pBS101). Appl. Biochem. Microbiol. 35:375–379. Mergaert, J., and J. Swings. 1996. Biodiversity of microorganisms that degrade bacterial and synthetic polyesters. J. Indust. Microbiol. 17:463–469. Meyer, O., E. Stackebrandt, and G. Auling. 1993. Reclassification of ubiquinone Q-10 containing carboxidotrophic bacteria: transfer of “[Pseudomonas] carboxydovorans” OM5T to Oligotropha, gen. nov., as Oligotropha carboxydovorans, comb. nov., transfer of “[Alcaligenes] carboxydus” DSM 1086T to Carbophilus, gen. nov., as Carbophilus carboxidus, comb. nov., transfer of “[Pseudomonas] compransoris” DSM 1231T to Zavarzinia, gen. nov., as Zavarzinia compransoris, comb. nov., and amended descriptions of the new genera. Syst. Appl. Microbiol. 16:390–395. Moebus, E., M. Jahn, R. Schmid, D. Jahn, and E. Maser. 1997. Testosterone-regulated expression of enzymes involved in steroid and aromatic hydrocarbon catabolism in Comamonas testosteroni. J. Bacteriol. 179:5951–5955. Morgan, C. A., and R. C. Wyndham. 1996. Isolation and characterization of resin acid degrading bacteria found

CHAPTER 3.2.7 in effluent from a bleached kraft pulp mill. Can. J. Microbiol. 42:423–430. Mudd, S., and S. Warren. 1923. A readily cultivable vibrio, filterable through Berkefeld “V” candles, Vibrio percolans (new species). J. Bacteriol. 8:447–458. Nozhevnikova, A. N., and G. A. Zavarzin. 1974. On the taxonomy of CO-oxidizing Gram-negative bacteria. Izv. Akad. Nauk SSSR, Ser. Biolog. 3:436–440. Oppenberg, H. R. S., G. Chen, T. Leisinger, and A. M. Cook. 1995. Regulation of the degradative pathways from 4toluenesulphonate and 4-toluenecarboxylate to protocatechuate in Comamonas testosteroni T-2. Microbiology 141:1891–1899. Oppermann, C. T., and E. Maser. 1996a. Characterization of a 3-α-hydroxysteroid dehydrogenase/carbonyl reductase from the Gram-negative bacterium Comamonas testosteroni. Eur. J. Biochem. 241:744–749. Oppermann, U. C. T., I. Belai, and E. Maser. 1996b. Antibiotic resistance and enhanced insecticide catabolism as consequence of steroid induction in the Gram-negative bacterium Comamonas testosteroni. J. Steroid Biochem. Molec. Biol. 58:217–223. Oppermann, U. C. T., C. Filling, K. D. Berndt, B, Persson, J. Benach, R. Ladenstein, and H. Joernvall. 1997. Active site directed mutagenesis of 3β/17β-hydroxysteroid dehydrogenase establishes differential effects on shortchain dehydrogenase/reductase reactions. Biochemistry (Wash.) 36:34–40. Palleroni, N. J. 1984. Genus I. Pseudomonas Migula 1894. In: N. R. Krieg and J. G. Holt (ed.) Bergey’s manual of systematic bacteriology. Williams and Wilkins. Baltimore, MD. 1:141–199. Parales, R. E., T. A. Ontl, and D. T. Gibson. 1997. Cloning and sequence analysis of a catechol 2,3-dioxygenase gene from the nitrobenzene-degrading strain Comamonas sp. JS765. J. Indust. Microbiol. Biotechnol. 19:385– 391. Patureau, D., N. Bernet, and R. Moletta. 1997. Combined nitrification and denitrification in a single aerated reactor using the aerobic denitrifier Comamonas sp. strain SGLY2. Water Res. 31:1363–1370. Petruschky, J. 1896. Bacillus faecalis alcaligenes (n. sp.). Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. I 19:187–191. Pickett, M. J., and J. R. Greenwood. 1986. Pseudomonas alcaligenes and Pseudomonas testosteroni: characterization and identification. Curr. Microbiol. 13:197–201. Proksova, M., D. Sladekova, B. Uhnakova, J. Augustin, and D. Toth. 1999. Primary biodegradtion and assimilation of dialkyl sulphosuccinate by Comamonas terrigena N3H. Biologia 54:671–677. Quinteros, R., S. Goodwin, R. W. Lenz, and W. H. Park. 1999. Extacellular degradation of medium chain length poly(beta-hydroxyalkanoates) by Comamonas sp. Int. J. Biol. Macromol. 25:135–143. Rubin, S. J., P. A. Granato, and B. L. Wasilauskas. 1985. Glucose-nonfermenting Gram-negative bacteria. In: Lennette, Balows, Hausler and Shadomy (Eds.) Manual of clinical microbiology, 4th Ed. American Society for Microbiology. Washington, DC. 330–349. Schach, S., B. Tshisuaka, S. Fetzner, and F. Lingens. 1995. Quinoline 2-oxidoreductase and 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase from Comamonas testosteroni 63: the first two enzymes in quinoline and 3-methylquinoline degradation. Eur. J. Biochem. 232: 536–544.

Comamonas

735

Schlaefi, H. R., M. A. Weiss, T. Leisinger, and A. M. Cook. 1994. Terephthalate 1,2-dioxygenase system from Comamonas testosteroni T-2: Purification and some properties of the oxygenase component. J. Bacteriol. 176:6644– 6652. Shinomiya, M., T. Iwata, K. Kasuya, and Y. Doi. 1997. Cloning of the gene for poly(3-hydroxybutyric acid) depolymerase of Comamonas testosteroni and functional analysis of its substrate-binding domain. FEMS Microbiol. Lett. 154:89–94. Skerman, V. B. D., V. McGowan, and P. H. A. Sneath. 1980. Approved lists of bacterial names. Int. J. Syst. Bacteriol. 30:225–420. Springael, D., J. van Thor, H. Goorissen, A. Ryngaert, R. De Baere, P. Van Hauwe, L. C. M. Commandeur, J. R. Parsons, R. De Wachter, and M. Mergeay. 1996. RP4::Mu3A-mediated in vivo cloning and transfer of a chlorobiphenyl catabolic pathway. Microbiology 142: 3283–3293. Stanier, R. Y., N. J. Palleroni, and M. Doudoroff. 1966. The aerobic pseudomonads: A taxonomic study. J. Gen. Microbiol. 43:159–271. Stigter, E. C. A., G. A. H. De Jong, J. A. Jongejan, J. A. Duine, J. P. Van Der Lugt, and W. A. C. Somers. 1997. Electron transfer and stability of a quinohaemoprotein alcohol dehydrogenase electrode. J. Chem. Technol. Biotechnol. 68:110–116. Stoorvogel, J., D. E. Kraayveld, C. A. Van Sluis, J. A. Jongejan, S. De Vries, and J. A. Duine. 1996. Characterization of the gene encoding quinohaemoprotein ethanol dehydrogenase of Comamonas testosteroni. Eur. J. Biochem. 235:690–698. Stoppel, R. D., and H. G. Schlegel. 1995. Nickel-resistant bacteria from anthropogenically nickel-polluted and naturally nickel-percolated ecosystems. Appl. Environ. Microbiol. 61:2276–2285. Sylvestre, M., Y. Hurtubise, D. Barriault, J. Bergeron, and D. Ahmad. 1996a. Characterization of active recombinant 2,3-dihydro-2,3-dihydroxybiphenyl dehydrogenase from Comamonas testosteroni B-356 and sequence of the encoding gene (bphB). Appl. Environ. Microbiol. 62:2710–2715. Sylvestre, M., M. Sirois, Y. Hurtubise, J. Bergeron, D. Ahmad, F. Shareck, D. Barriault, I. Guillemette, and J. M. Juteau. 1996b. Sequencing of Comamonas testosteroni strain B356 biphenyl/chlorobiphenyl deoxygenase genes: Evolutionary relationships among Gram-negative bacterial biphenyl dioxygenases. Gene 174:195–202. Takahashi, E., M. Furui, and T. Shibatani. 1997. Scale-up of D-lysine production from L-lysine by successive chemical racemization and microbial asymmetric degradation. Biotechnol. Lett. 19:245–249. Talalay, P., M. Mollomo Dobson, and D. F. Tapley. 1952. Oxidative degradation of testosterone by adaptive enzymes. Nature (London) 170:620–621. Tamaoka, J., D.-M. Ha, and K. Komagata. 1987. Reclassification of Pseudomonas acidovorans den Dooren de Jong 1926 and Pseudomonas testosteroni Marcus and Talalay 1956 as Comamonas acidovorans comb. nov. and Comamonas testosteroni comb. nov., with an emended description of the genus Comamonas. Int. J. Syst. Bacteriol. 37:52–59. Tazuke, Y., K. Matsuda, K. Adachi, and Y. Tsukada. 1994. Purification and properties of bile acid sulfate sulfatase from Pseudomonas testosteroni. Biosci. Biotechnol. Biochem. 58:889–894.

736

A. Willems and P. De Vos

Teramoto, M., H. Futamata, S. Harayama, and K. Watanabe. 1999. Characterization of a high-affinity phenol hydroxylase from Comamonas testosteroni R5 by gene cloning, and expression in Pseudomonas aeruginosa PAO1c. Molec. Gen. Genet. 262:552–558. Toth, D., J. Huska, I. Zavadska, and M. Dobrotova. 1996. Effect of bacterial starvation on surfactant biotransformation. Folia Microbiol. 41:477–479. Vanbrabant, J., P. De Vos, M. Vancanneyt, J. Liessens, W. Verstraete, and K. Kersters. 1993. Isolation and identification of autotrophic and heterotrophic bacteria from an autohydrogenotrophic pilot-plant for denitrification of drinking-water. Syst. Appl. Microbiol. 16:471–482. Vaneechoutte, M., R. Rossau, P. De Vos, M. Gillis, D. Janssens, N. Paepe, A. De Rouck, T. Fiers, G. Claeys, and K. Kersters. 1992. Rapid identification of bacteria of the Comamonadaceae with amplified ribosomal DNArestriction analysis (ARDRA). FEMS Microbiol. Lett. 93:227–234. Wen, A, M. Fegan, C. Hayward, and S. Chakraborty, and L. I. Sly. 1999. Phylogenetic relationships among members of the Comamonadaceae, and description of Delftia acidovorans (den Dooren de Jong 1926, and Tamaoka et al. 1987) gen. nov., comb. nov. Int. J. Syst. Bacteriol. 49:567–576. Willems, A., B. Pot, E. Falsen, P. Vandamme, M. Gillis, K. Kersters, and J. De Ley. 1991. Polyphasic taxonomic study of the emended genus Comamonas: Relationship

CHAPTER 3.2.7 to Aquaspirillum aquaticum, E. Falsen group 10, and other clinical isolates. Int. J. Syst. Bacteriol. 41:427–444. Xiong, G. M., H. J. Martin, C. Schafers, and E. Maser. 2001. A model on the regulation of 3 alpha-hydroxysteroid dehydrogenase/carbonyl reductase expression in Comamonas testosteroni. Chemico-biol. Interact. 13:723–736. Yap, L. F., Y. K. Lee, and C. L. Poh. 1999. Mechanism for phenol tolerance in phenol-degrading Comamonas testosteroni strains. Appl. Microbiol. Biotechnol. 51:833– 840. Yasuda, M., T. Sakamoto, R. Sashida, M. Ueda, Y. Morimoto, and T. Nagasawa. 1995. Microbial hydroxylation of 3cyanopyridine to 3-cyano-6-hydroxypyridine. Biosci. Biotechnol. Biochem. 59:572–575. Zhao, Q., Abeygunawardana, C., Talalay, P., and Mildvan, A. S. 1996. NMR evidence for the participation of a lowbarrier hydrogen bond in the mechanism of d5-3ketosteroid isomerase. Proc. Natl. Acad. Sci. 93:8220– 8224. Zhao, Q., Abeygunawardana, C., and Mildvan, A. S. 1997. NMR studies of secondary structure in isolation and the steroid binding site of d5-3-ketosteroid isomerase in complexes with diamagnetic and paramagnetic steroids. Biochemistry (Wash.) 36:3458–3472. Zhao, J. S., and O. P. Ward. 1999. Microbial degradation of nitrobenzene and mono-nitrophenol by bacteria enriched from municipal activated sludge. Can. J. Microbiol. 45:427–432.

Prokaryotes (2006) 5:737–746 DOI: 10.1007/0-387-30745-1_32

CHAPTER 3.2.8 ehT

areneG

mu i re t cabomorhC

dna

mu i re t cabon i htna J

The Genera Chromobacterium and Janthinobacterium MONIQUE GILLIS AND JOZEF DE LEY

Purple-pigmented bacteria have been described since the end of the 19th century; they were reported as discoloring a variety of natural materials and occasionally as the causative agent of septicemia in humans and animals. Bacteria producing purple and violet colonies due to the production of a nondiffusible pigment, violacein, were classified in a redefined genus Chromobacterium by Buchanan (1918). The structure of violacein, an indole derivative produced via the oxidation of tryptophan, is shown in Fig. 1. It can be easily identified (Johnson and Beer, 1971) spectrophotometrically from the following properties: 1) in ethanolic solution it has an absorption maximum at 579 nm and a minimum at 430 nm; 2) by adding 10% (v/v) H2SO4 the solution turns green with an absorption maximum at 700 nm; and 3) when NaOH is added to an ethanolic solution, the solution turns green and afterwards reddish brown. Recently, violacein has also been characterized by nuclear magnetic resonance spectroscopy and mass spectrometry (Riveros et al., 1988). The pigment is only abundantly produced when tryptophan is available in the culture medium. Although violacein is produced by only a few groups of bacteria, its presence does not necessarily indicate a close relationship between these organisms. An extensive phenotypic study (Sneath, 1956, 1960, 1974) within Chromobacterium, as defined by Buchanan (1918), provided evidence of two groups within this genus: a fermentative and mesophilic one (growth at 37°C but not at 4°C) and a nonfermentative and psychrophilic one (growth at 4°C but not at 37°C), for which he proposed two species, Chromobacterium violaceum and [Chromobacterium] lividum, respectively. However, these two species have been reported as being not closely related (Moffet and Colwell, 1968; Sneath, 1974) and since the latter was misnamed, in this chapter its epithet and all other misnamed epithets shall be enclosed in square brackets. Violacein is not a good taxonomic marker as shown by the fact that not only have nonpigmented Chromobacterium violaceum and [Chromobacterium] lividum strains been isolated, but also since marine, violacein-producing strains were isolated, which were assigned on phenotypic grounds to the genus Alteromonas (Gauthier, 1976, 1982). Later, Van Landschoot and De Ley (1983) confirmed that these marine strains were genuine Alteromonas. The relationships between the violacein-producing bacteria classified in Chromobacterium (Sneath, 1974) could finally be elucidated by De Ley et al. (1978). DNA-rRNA hybridizations within and between the violaceum and the lividum taxons and with different other Gram-negative genera demonstrated clearly that these purple-pigmented bacteria represent two tight but separate groups within rRNA This chapter was taken unchanged from the second edition.

superfamily III (see also De Ley’s introduction to the Proteobacteria in The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition) which are less related to each other than to other genera from this rRNA superfamily. In consequence, a new genus Janthinobacterium with one species J. lividum was created for the lividum taxon and the genus name Chromobacterium was restricted to the mesophilic, fermentative violaceum taxon (De Ley et al., 1978). Chromobacterium constitutes a separate rRNA branch on which all violaceum strains cluster with a Tm(e) of 76.5 to 80°C. Janthinobacterium constitutes a separate rRNA subbranch which is closely related to the rRNA subbranch containing [Pseudomonas] rubrisubalbicans and a yet unnamed group of bacteria isolated from clinical material (see Fig. 6 in The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition). The Janthinobacterium-[Pseudomonas] rubrisubalbicans rRNA branch is more closely related to the solanacearum rRNA branch than to Chromobacterium, emphasizing the deep phylogenetic gap between Chromobacterium and Janthinobacterium. Cataloging results (Woese et al., 1984) confirm the separate position of both genera in the beta group (Stackebrandt et al., 1988). In this chapter, Chromobacterium and Janthinobacterium are treated together merely on historical grounds. Moss et al. 1978 described a new group of violaceincontaining, fermentative, psychrophilic, fresh water bacteria and assigned them to a new species fluviatile in the genus Chromobacterium. It differs from C. violaceum by a thin flat, spreading colony type, some other phenotypic features (see below) and % GC values of 50 to 52. Within C. violaceum the GC content varies from 65 to 68 mol%, indicating at once that these new organisms cannot be members of Chromobacterium. DNA-rRNA hybridizations (Moss and Bryant, 1982) demonstrated that [Chromobacterium]* fluviatile is not closely related to the C. violaceum rRNA cluster. With a Tm(e) of 70 to 71°C vs. the C. violaceum rRNA probe, they occupy a separate position and are further removed from C. violaceum than the latter species is from the Neisseriaceae (Rossau et al., 1989). Comparison of their 16S rRNA sequences corroborates this conclusion (Dewhirst et al., 1989). Therefore, these organisms cannot belong in Chromobacterium; they have thus been misnamed and shall not be discussed here further. The differentiation of these three groups of violacein-producing organisms is given in Table 1.

*Since this manuscript was submitted, Logan (1989) proposed a monospecific genus Iodobacter to accomodate the [Chromobacterium] fluviatile group. Their phenotypic results on Iodobacter and the “atypical, Janthinobacterium lividum strains” are integrated in Table 1.

738

M. Gillis and J. de Ley

CHAPTER 3.2.8

Chromobacterium The following description of the genus is based on that of Chromobacterium violaceum by Sneath (1974, 1984). The genus Chromobacterium consists of Gram-negative, oxidase-positive, catalase-positive rods with rounded ends which are usually motile by both a single polar flagellum and by one to four lateral flagella. The two types of flagella differ in their wave types, staining capacity, and antigenic characteristics. Chromobacteria are facultative anaerobes and produce violet colonies on solid media; a violet ring is formed in liquid media at the surface with a fragile pellicle. Growth is best at 30 to 35°C; the minimum growth temperature is 10 to 15°C and the maximum is 40°C (20% of the strains can grow at 44°C). They are chemoorganotrophs

using mainly carbohydrates fermentatively. Nitrate and usually nitrite are reduced, mostly with a variable gas production. They grow on ordinary media and are resistant to benzylpenicillin (10 µg/ml) and to vibriostatic agent 0/129 [2,4-diamino-6,7-diisopropylpteridine (30 µg/ disc)]. The GC content of the DNA is 65 to 68 mol%. Other characteristics are as described by Sneath (1984) for C. violaceum. The type and sole species is C. violaceum. Between representative strains of this species, more than 83% DNA binding was found (De Ley et al., 1978). C. violaceum is a saprophyte from soil and water and is normally considered as nonpathogenic for humans. Occasionally, however, it is an opportunistic pathogen of extreme virulence for humans and animals (see below).

Habitats H N

C. violaceum is mainly found in soil and water (Sneath, 1956), where it usually constitutes only a minor component of the total microflora. It is found in temperate but more frequently in tropical and subtropical regions. It was suggested that it can be isolated more frequently from soil than from water (Corpe, 1951; Moss and Ryall, 1981). Its role in the rhizosphere of plants is not clear, although Hussain and Van cˇ ura (1970) isolated C. violaceum strains from rhizosphere soil of maize and demonstrated that the inoculation of maize

O

HO

N H

O

N H

Fig. 1. Structure of violacein.

Table 1. Characteristics differentiating the groups of violacein-producing organisms. Character Growth at 4°C Growth at 37°C Fermentation of glucose (O/F test) Anaerobic growth Cyanide produced Turbid zone on egg yolk agar (lecithinase) Acid from Trehalose L-arabinose D-cellobiose D-galactose Gluconate D-maltose N-acetylglucosamine Lactate utilization Casein hydrolysis Esculin hydrolysis Arginine decarboxylase GC content (mol%) a

Chromobacterium

Janthinobacterium

Iodobacter [Chromobacterium] fluviatilis

− + Fb + + +

+ (96%)a − 0c (90%) − (96%) − −

+ − F + − (+)d

+ − − − + − + + + − + 65–68

− (82%) + + (99%) + (96%) − (99%) + (96%) − (99%) + (99%) − or (+) + (80%) − 65–66

+ − − − + (96%) + + − + − − 50–52

Numbers in parentheses indicate the % of strains giving the indicated reaction. F, fermented. c O, oxidized. d (+) weakly positive. The phenotypic data are from Sneath (1984), Moss et al. (1978) and Logan (1989). The GC values are from De Ley et al. (1978) and from Moss et al. (1978) for Iodobacter [Chromobacterium] fluviatilis. b

CHAPTER 3.2.8

The Genera Chromobacterium and Janthinobacterium

seeds with such C. violaceum strains significantly increased the yield of dry matter of the plant. C. violaceum could not be recovered from food products (Koburger and May, 1982). C. violaceum has been isolated from severe infections in humans and animals; water or soil contaminated with C. violaceum is mostly the causal agent. Although a C. violaceum infection in humans can result in mild diarrhea with a subsequent recovery (Sneath et al., 1953), it is mostly fatal for both humans and animals. A review of the earlier literature concerning chromobacteriosis was given by Sneath (1960). Later rare fatal cases of infection with C. violaceum were reported in different countries, i.e., the United States (Johnson et al., 1971), Vietnam (Ognibene and Thomas, 1970), Cuba (Machin Villafranca et al., 1986), Taiwan (Wu et al., 1986), Argentina (Kaufman et al., 1986), Nigeria (Onile et al., 1984), Brazil (Petrillo et al., 1984), and Australia (Wilkey and McDonald, 1983). The symptoms of the infection can vary considerably but lead mostly to septicemia, causing a rapid death. In the different case reports the symptoms were liver and lung abscesses (Sneath, 1960), meningitis (Shetty et al., 1987), adenitis (Sorensen et al., 1985), periorbital cellulitis (Simo et al., 1984; Feldman et al., 1984), toxemia, and multiple skin abscesses (Petrillo et al., 1984). Only exceptionally nonfatal chromobacterial sepsis has been reported as, i.e., a case of multiple liver abscesses (Suarez et al., 1986) or a systemic chromobacteriosis (Victorica et al., 1974). The disease can be cured by tetracyclines if treated at an early stage (Moss and Ryall, 1981). A recent comparison of the in vitro activity of 25 antimicrobial agents against clinical strains of C. violaceum showed that ciprofloxacin, norfloxacin, and perfloxacin are also highly active (Aldridge et al., 1988). For some eye infections, early recognition and aggressive treatment (with tetracycline, chloramphenicol, and cotrinoxazole) may result in a higher cure rate (Feldman et al., 1984). Infections in animals were also described and studied intensively, i.e., in swine (Laws and Hall, 1964; Liu et al., 1988), Malayan gibbons, a Malayan sun bear (Groves et al., 1969), an Assam macaque (McClure and Chang, 1976), a dog (Gogolewski, 1983), and monkeys (Collins et al., 1985). Virulence for mice and the pathology of infections appear to be similar for both nonpigmented and pigmented C. violaceum strains (Sivendra and Tan, 1977). Virulent C. violaceum strains produce an endotoxin which is more reactive than that from avirulent strains and it is suggested that the virulent strains are protected from phagocytic attack by elevated levels of superoxide dismutase and catalase (Miller et al.,

739

1988). Experimental infections with C. violaceum or with its endotoxin were studied in mice and pigs (Zwang et al., 1987; Liu et al., 1988).

Enrichment and Isolation The medium and conditions used to isolate and enumerate C. violaceum can strongly influence their recovery (Ryall and Moss, 1975; Koburger and May, 1982). Chromobacteria can grow easily on common laboratory media such as nutrient agar or GYCA (1% glucose, 0.5% yeast extract, 3% CaCO3, and 2% agar in distilled water), and violacein is produced in the presence of oxygen. Sneath (1960) concluded that blood agar base was more suitable for the isolation of small numbers of C. violaceum from medical sources, because they can be inhibited by the presence of peroxide which can be present in nutrient agar. For isolation from soil and water, the common media are not so suitable because chromobacteria represent only a minor part of the present microflora, so that large numbers of other bacteria can overgrow or prevent chromobacteria from growing well and producing violacein, which is an important feature in screening. Therefore, an enrichment procedure or a selective medium is recommended. In most cases, the selective media enrich for Chromobacterium as well as for Janthinobacterium and [Chromobacterium] fluviatile. The latter two taxa can be differentiated from C. violaceum afterwards by their phenotypic features (see Table 1). Corpe (1951) described the following simple enrichment technique: “Five grams samples of soil were placed in sterile Petri dishes and soaked with sterile distilled water. Sterile polished or precooked rice grains were sprinkled over the surface of the soil and the plates incubated at 23– 25°C for five days. At the end of the incubation period, the rice grains were partially covered with purple membranous growth indicating the presence of Chromobacterium sp.” Ryall and Moss (1975) used a selective medium based on quarter strength nutrient agar to count chromobacteria and janthinobacteria in water samples: Composition of the Medium of Ryall and Moss (1975). Beef extract (Oxoid) Yeast extract Peptone NaCl Agar Distilled water

0.25 g 0.5 g 1.25 g 1.25 g 20 g 1 liter

To the molten agar medium (45°C), filter-sterilized solutions of colistin and sodium deoxycholate were added to give final concentrations of 15 µg/ml and 0.3 mg/ml, respectively. For counting chromobacteria and janthinobacteria in soil samples, cycloheximide was also added to

740

M. Gillis and J. de Ley

give a final concentration of 30 µg/ml to prevent overgrowth by fungi.

Composition of the Medium of Keeble and Cross (1977). An improved selective medium (Keeble and Cross, 1977) is based on Bennett’s agar: Yeast extract (Difco) Beef extract (Oxoid) Bacto-casitone (Difco) Glucose Agar (Lab. M no. 2) Distilled water

1g 1g 2g 10 g 18 g 1 liter

To the molten agar medium were added filter-sterilized solutions of neomycin hydrochloride, cycloheximide, and nystatin to give a final concentration of 50 µg/ml of each. This medium is described as improving the pigmentation.

After comparing several media, Koburger and May (1982) reported that Bennett’s agar was best for the isolation of both C. violaceum and Janthinobacterium lividum when incubated at 25°C but that at 35°C Aeromonas membrane agar gave the highest recoveries of C. violaceum. Aeromonas membrane agar (Rippey and Cabelli, 1979) is prepared as follows: Tryptose Trehalose Yeast extract NaCl KCl MgSO4 ·7H 2O FeCl3 ·6H 2O Bromothymol blue Deionized water

0.5 g 0.5 g 0.2 g 0.3 g 0.2 g 0.02 g 0.01 g 0.004 g 100 ml

The ingredients are dissolved at room temperature, the pH is adjusted to 8.0 with 10 N NaOH, 1.5 g agar is added, and the mixture is autoclaved at 121°C for 15 min. After the addition of 1.0 ml ethanol, the mixture is cooled to 50°C, and 2 mg ampicillin and 10 mg desoxycholate are added. It is recommended that the poured plates be stored in the dark at 4°C and used within 6 weeks.

For inoculation, 0.1-ml samples are pipetted onto the surface and spread. They are allowed to dry for 30 min and incubated at 25 and 35°C for 7 and 5 days, respectively. All purple colonies developing on the plate were isolated and identified. Koburger and May (1982) suggested that the failure in other studies (Ryall and Moss, 1975; Keeble and Cross, 1977) to recover C. violaceum from soil and water and their own failure to isolate it from a limited number of food products can be related to the temperature sensitivity of these organisms. C. violaceum dies rapidly at 4°C. Chromobacterium can also be isolated on a citrate-ammonium salts medium (Moss and Ryall, 1981) as described for Janthinobacterium further in this chapter. Preservation of Strains The organisms can survive for several years in dilute peptone water (0.1% peptone) at room temperature. On

CHAPTER 3.2.8

GYCA or nutrient agar slants they can be kept for some months. Lyophilization and L-drying can be used to preserve them for longer periods. Freezing in nutrient broth containing 15% glycerol with storing at −80°C or in liquid N2 is also recommended.

Identification When definitely pigmented, Chromobacterium has only to be differentiated from other violacein-producing nonmarine strains belonging in the genus Janthinobacterium or to [Chromobacterium] fluviatile. The differentiating features are given in Table 1. Chromobacterium differs from Alteromonas luteoviolacea in its higher GC content and its ability to use citrate, glucose, fructose, and mannose (Gauthier, 1976, 1982). Nonpigmented C. violaceum strains can be confused with members of Vibrio and Aeromonas. However, C. violaceum differs from Aeromonas in its flagellar morphology, its negative reaction for indole and for methyl red, and its production of HCN. Moreover, C. violaceum differs from Vibrio in its GC content and by a negative response to the lysine and ornithine decarboxylase tests but by a positive reaction in the arginine decarboxylase test. Unlike C. violaceum, most strains of the genera Aeromonas and Vibrio are able to use D-mannitol as a carbon source (Sneath, 1984; Sivendra and Tan, 1977). Singh et al. (1988) used a DNA dot blot technique and a DNA probe from their nonpigmented isolate to differentiate the latter strain from Aeromonas and to identify it as C. violaceum. Members of Chromobacterium can easily be identified by hybridization with a 3H-labelled 23S rRNA probe from C. violaceum NCTC 9757: only members of C. violaceum have Tm(e) values between 76.5 and 80°C. [Chromobacterium] fluviatile strains have Tm(e) values of 70 to 71°C. Members of Janthinobacterium have Tm(e) values of 67 to 68°C and can further be identified by hybridization with a 3H-labelled 23S rRNA probe of J. lividum NCTC 9796. All other taxa are below 72°C Tm(e).

Special Features and Applications In this part, we call attention to products or metabolic pathways that have been intensively studied in C. violaceum and their possible applications. First of all, the biosynthesis of violacein (Riveros et al., 1988) and the different effects of this pigment have been studied. Violacein has antibiotic activity (DeMoss, 1967; Hoshino et al., 1987) and protects against predation by protozoa. In the older literature (Burbanck, 1942; Singh,

CHAPTER 3.2.8

The Genera Chromobacterium and Janthinobacterium

1942, 1945), C. violaceum and violacein itself are described as lethal to some ciliated protozoa. Amoebae and flagellates can also be killed by these bacteria (Singh, 1942). Later Groscop and Brent (1964) showed that violacein is toxic to soil amoebae while C. violaceum is toxic to Vorticella microstoma (Curds and Vandyke, 1966). Violacein has also a trypanocide effect (Riveros et al., 1988). The production of violacein can be used as a microbiological assay for L-tryptophan (Sebek, 1965). Apart from violacein, C. violaceum produces other antibiotics. Several were isolated and characterized in the last years: aerocyanidin, bearing an isonitrile group and being active against Gram-positive bacteria (Parker et al., 1988); aerocavin, exhibiting in vitro activity against both Gram-negative and Gram-positive bacteria (Singh et al., 1988); 3,6-dihydroxyindoxazene, inhibiting growth of Gram-negative bacteria (Hamada et al., 1983); and factor Y-T0678H (6hydroxy-3-oxo-1,2-benzisoxazolin), exhibiting a selective activity against Gram-negative bacteria (Imai et al., 1983). Other important compounds are produced by C. violaceum: arphamenines A and B, two specific inhibitors of aminopeptidase B that enhance immune responses (Umezawa et al., 1983). The production and function of C. violaceum aminopeptidases involved in enkephalin hydrolysis was studied by Weiss et al. (1983). Cooper et al. (1985) studied novel potentiators of β-lactam antibiotics and isolated two new compounds, SQ 28,504 and SQ 28,546, from a C. violaceum strain. These novel bacterial metabolites are glycopeptides which act synergistically with β-lactam antibiotics against Gram-negative bacteria. Earlier β-lactamase, primarily active against cephalosporins, was shown in a C. violaceum strain (Farrar and O’Dell, 1976). C. violaceum has also been used in studies on the production of HCN (Brysk et al., 1969; Niven et al., 1975), of unusual sugar compounds (Stevens et al., 1963), and of extracellular polysaccharides (Corpe, 1964). The tough membranous aspect of some C. violaceum colonies is the result of the latter polysaccharides (Corpe, 1964). Their role in the formation of soil aggregates has been suggested (Martin and Richards, 1963). The Copper-containing phenylalanine monooxygenase (Benkovic et al., 1986; Pember et al., 1987a, 1987b; Fujisawa and Nakata, 1987) and the denitrification processes (Bazylinski et al., 1986) were intensively studied. It was shown that denitrification terminates with the production of N2O and that NO2− reduction to N2O is not coupled to growth but may serve as a detoxification mechanism.

741

A C. violaceum strain has been isolated that can grow on and use acetonitrile as sole carbon source (K. D. Chapatwale et al., 1988) Another C. violaceum strain solubilized a considerable amount of pure gold after 40 days incubation in nutrient medium because the cyanide produced during growth and stationary phase formed the complex anion [Au(CN)2]− with gold (Smith and Hunt, 1985).

Janthinobacterium The basic description of this genus was given by Sneath in 1984. Janthinobacterium consists of Gram-negative rods with rounded ends, sometimes slightly curved. No resting stages are formed. They are usually motile by means of both a single polar flagellum and by one to four lateral flagella. They are strict aerobes. They produce violet colonies on solid media; in nutrient broth a violet ring is formed at the junction of the liquid surface and the container wall. They are resistant to benzylpenicillin (10 µg/ml) and to vibriostatic agent 0/129 (30 µg/disc). The most striking features are: 1) janthinobacteria are aerobic chemoorganotrophs with a strictly respiratory type of metabolism with oxygen as the terminal electron acceptor. 2) They have an optimum growth temperature of 25°C, a maximum growth temperature of 32°C and a minimum growth temperature of 2°C. 3) Only a few strains produce HCN. 4) They can grow rapidly on a medium containing citrate and ammonia as sole carbon and nitrogen source. 5) Nitrate and nitrite are reduced mostly with visible gas production. 6) The GC content is 61 to 67 mol%. They are soil and water organisms (Moss et al., 1978; Koburger and May, 1982) common in temperate climates. They are sometimes the causative agent of food spoilage. The type and sole species is J. lividum. Within this species, percentages of DNA binding of at least 48% were encountered between representative strains (De Ley et al., 1978). The membraneforming strains, formerly named C. amethystinum or C. membranaceum and sometimes referred to as “atypical C. lividum strains” (Moss and Ryall, 1981), and an unusual strain (ATCC 14487) which preferentially grows at 4°C all belong in J. lividum (De Ley et al., 1978; Sneath, 1984). The typical features in which Janthinobacterium differs from Chromobacterium can be found in Table 1.

Habitats J. lividum strains have been isolated from water and soil from rivers, creeks, lakes, and springs (Ryall and Moss, 1975; Koburger and May, 1982; Williams and Albright, 1984; Quevedo-

742

M. Gillis and J. de Ley

Sarmiento et al., 1986; Gonzalez et al., 1987). They mostly comprise only a small percentage of the total microflora, e.g., a mean of 5% of the total aerobic heterotrophic bacterial population in natural spring water (Quevedo-Sarmiento et al., 1986); a mean of 915 J. lividum isolates in 1 g water or soil (Koburger and May, 1982); and less than 1 to 3% of the total population sampled in lake water (Williams and Albright, 1984). According to Moss et al. (1978) they constitute approximately 75% of the total amount of purple-pigmented bacteria isolated from water and soil samples in a lowland river (river Wey in the UK). They are frequently isolated from a variety of foods, i.e., processed poultry (Cox, 1975); turbot (Mudarris and Austin, 1988); various vegetables (Koburger and May, 1982); and Japanese noodles (Naito et al., 1986a). In turbot gills (Mudarris and Austin, 1988), J. lividum represented approximately one-third of the aerobic heterotrophic bacteria (7 × 105/g wet weight of gill tissue). In vegetables, they are present in quantities varying from 130 to over 6000/g (Koburger and May, 1982). Isolates from slaughterhouse effluents (Etherington et al., 1976) were colorless, could grow on meat, and exhibited a high proteolytic activity (Dainty et al., 1978), which is exceptional for members of this genus (Sneath, 1984). Bettelheim et al. (1968) isolated bacterial strains from germinating seeds of Psychotria nairobiensis (Rubiaceae) and Ardisia crispa (Myrsinaceae), which they identified then as C. lividum, thus now J. lividum. By an immunofluorescence technique, Bettelheim et al. (1968) detected these organisms in tissues of several leaf-nodulated Rubiaceae and Myrsinaceae, and presented the hypothesis that some J. lividum strains are bacterial endophytes. We shall not review the discussions and disagreements in the literature on this issue (Lersten and Horner, 1976; see also The Genera Chromobacterium and Janthinobacterium in Volume 5) because it was shown that Bettelheim’s isolates are not genuine janthinobacteria (De Ley et al., 1978) since they belong in rRNA superfamily IV, where they are a member of the [Flavobacterium] capsulatum rRNA branch (The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition). The role, if any, of J. lividum strains in the induction of sporangia in Phytophthora cinnamomi has been discussed (Zentmeyer, 1965; Broadbent et al., 1974).

Enrichment and Isolation As mentioned before, Janthinobacterium can be enriched and isolated on the same media as given in detail for Chromobacterium above.

CHAPTER 3.2.8

According to Moss and Ryall (1981), they grow rapidly and can be selected on a citrate ammonium salts agar medium: Citric acid NaCl MgSO4 ·7H 2O NH4H2PO4 K2HPO4 Ionoagar nr. 2 (Oxoid) Distilled water

2g 1g 0.2 g 1g 1g 15 g 1 liter

Koburger and May (1982) recovered J. lividum from a variety of food samples only on Bennett’s agar (see above) at 25°C. On this medium they found higher counts in water and soil samples than on the medium of Ryall and Moss (1975) or on Aeromonas membrane agar (Rippey and Cabelli, 1979). Maximal numbers of J. lividum were isolated at 25°C from turbot gills (Mudarris and Austin, 1988) on the following medium (%w/v or v/v): CaCl2 Yeast extract Beef extraxt Casein Tryptone Agar no. 1 Aged sea water pH

0.1 0.1 0.5 0.6 0.2 1.5 75 7.2

Identification Characteristics differentiating J. lividum from C. violaceum and Iodobacter fluviatilis* have been given in Table 1. Differences with Alteromonas luteoviolacea have been reported above. Although colorless J. lividum strains could be confused with Pseudomonas fragi (Moss and Ryall, 1981), Dainty et al. (1978) were able to identify correctly their nonpigmented isolates as J. lividum. Because J. lividum strains constitute a narrow rRNA cluster with Tm(e) values between 77 and 79°C vs. 3H labelled 23S rRNA from J. lividum NCTC 9796, they can easily be identified by hybridizing with this rRNA probe. [Pseudomonas] rubrisubalbicans, [Aquaspirillum] autotrophicum, and an as yet unnamed group of strains isolated from clinical material and described by E. Falsen (Culture Collection, University of Göteborg, Sweden) have Tm(e) values of approximately 76°C and are the closest neighbors of J. lividum (see The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition). Phenotypically (M. Goor et al. 1986; see The Genus Aquaspirillum in Volume 5) it is possible to differentiate J. *The latter species was originally named Iodobacter fluviatile, but since Iodobacter is masculine, it obviously has to be Iodobacter fluviatilis.

CHAPTER 3.2.8

The Genera Chromobacterium and Janthinobacterium

743

Table 2. Characteristics differentiating Janthinobacterium from [Pseudomonas] rubrisubalbicans and similar unnamed clinical isolates. Character Urease Hydrolysis of esculin Growth on D-fucose, 5-keto-gluconate, acetate, caprate, amylamine, and tryptamine Growth on L-histidine

Janthinobacterium

[Pseudomonas] rubrisubalbicans and unnamed clinical isolatesa

− + −

+ − +

+

−

Isolates of Falsen (1989).

lividum from the group formed by [Pseudomonas] rubrisubalbicans and the unnamed clinical isolates (Table 2) and from [Aquaspirillum] autotrophicum (see Table 1 in The Genus Aquaspirillum in Volume 5). Results from M. Goor et al. (1986). Preservation of Strains J. lividum strains survive for several years at 4°C in 0.1% peptone or at −80°C in nutrient broth with 15% glycerol as a cryoprotectant. Lyophilization and storage at 4°C is also applied although the viability may be low (Sneath, 1984).

Special Features and Applications For the different effects of violacein, we refer to the discussion under “Chromobacterium” above. The antimicrobial activity of violacein isolated from J. lividum has recently been described (Naito et al., 1986b). The location of the pigment was investigated by Lin and Kester (1988), and in broth cultures, both intracellular and extracellular pigment was found. Intracellular pigment is located in the cell membrane. Extracellular pigment is water soluble and shows a blue shift when compared to solvent-extracted pigment. It was suggested that extracellular pigment is possibly non-covalently bound to a small protein. J. lividum produces 2,5-diketogluconate (Bernaerts and De Ley, 1971) via 2-ketogluconate. Kwok et al. (1987) described a J. lividum strain as a bacterial antagonist inducing suppression of Rhizoctonia damping-off in container media amended with composted hardwood tree bark. Strains from food produce sometimes extracellular metallo-proteinases (Etherington et al., 1976; Dainty et al., 1978) which could play a role in tenderizing meat. A J. lividum strain was used to study the effects of the insecticide acephate on bacterial growth and nutrient uptake (Williams and Albright, 1984). Acknowledgments. We are indebted to the National Fund for Scientific Research (Belgium) and the Fund for Medical Scientific Research (Belgium) for research and personnel grants.

Literature Cited Aldridge, K. E., G. T. Valainis, C. V. Sanders. 1988. Comparison of the in-vitro activity of ciprofloxacin and 24 other antimicrobial agents against clinical strains of Chromobacterium violaceum. Diagn. Microbiol. Infect. Dis. 10:31– 40. Bazylinski, D. A., E. Palome, N. A. Blakemore, R. P. Blakemore. 1986. Denitrification by Chromobacterium violaceum. Appl. Environm. Microbiol. 52:696–699. Benkovic, S. J., L. M. Bloom, G. Bollag, T. A. Dix, B. J. Gaffney, S. Pember. 1986. The mechanism of action of phenylalanine hydroxylase. Ann. N.Y. Acad. Sci. 471:226–232. Bernaerts, M., J. De Ley. 1971. 2,5-Diketogluconate formation by Chromobacterium. J. Microbiol. Serol. 37:185– 195. Bettelheim, K. A., J. F. Gordon, J. Taylor. 1968. The detection of a strain of Chromobacterium lividum in the tissues of certain leaf nodulated plants by the immunofluorescence technique. J. Gen. Microbiol. 54:177–184. Broadbent, P., K. F. Baker, J. Aust. 1974. Association of bacteria with sporangium formation and breakdown of sporangia in Phytophthora sp. Austr. J. Agricultural Research 25:139–145. Brysk, M. M., W. A. Corpe, L. V. Hankes. 1969. β-Cyanoalanine formation by Chromobacterium violaceum. J. Bacteriol. 97:322–327. Buchanan, R. E. 1918. Studies in the nomenclature and classification of bacteria. J. Bacteriol. 3:27–61. Burbanck, W. D. 1942. Physiology of the ciliate Colpidium colpoda. Physiological Zoology 15:342–362. Chapatwala, K. D., J. D. Richardson, M. Nawaz, J. H. Wolfram. 1988. Isolation and identification of acetonitrile degrading bacteria. Abstr. Annual Meeting American Society Microbiology, May 8–13, Miami Beach, Florida, USA. 88:225. Collins, B., G. Kolias, A. Battles, A. Moreland. 1985. Chromobacterium violaceum infection in a stumptailed monkey Macaca arctoides. Lab. Anim. Sci. 35:530–531. Cooper, R., J. S. Wells, R. B. Sykes. 1985. Novel potentiators of β-lactam antibiotics. J. Antibiot. 38:449–454. Corpe, W. A. 1951. A study of the wide spread distribution of Chromobacterium species in soil by a simple technique. J. Bacteriol. 62:515–517. Corpe, W. A. 1964. Factors influencing growth and polysaccharide formation by strains of Chromobacterium violaceum. J. Bacteriol. 88:1433–1441. Cox, N. A. 1975. Isolation and identification of a genus, Chromobacterium, not previously found on processed poultry. Appl. Microbiol. 29:864.

744

M. Gillis and J. de Ley

Curds, C. R., J. M. Vandyke. 1966. The feeding habits and growth rates of some fresh-water ciliates found in activated sludge plants. J. Appl. Ecology 3:127–137. Dainty, R. H., D. J. Etherington, B. G. Shaw, J. Barlow, G. T. Banks. 1978. Studies on the production of extracellular proteinases by a non-pigmented strain of Chromobacterium lividum isolated from abattoir effluent. J. Appl. Bacteriol. 45:111–124. De Ley, J., P. Segers, M. Gillis. 1978. Intra-and intergeneric similarities of Chromobacterium and Janthinobacterium ribosomal ribonucleic acid cistrons. Int. J. Syst. Bacteriol. 28:154–168. DeMoss, R. D. 1967. Violacein. 77–81.D. Gottlieb, and P. Shaw (ed.) Mechanisms of action and biosynthesis of antibiotics, vol. 2. Springer-Verlag. New York. Dewhirst, F. E., B. J. Paster, P. L. Bright. 1989. Chromobacterium, Eikenella, Kingella, Neisseria, Simonsiella and Vitreoscilla species comprise a major branch of the beta group Proteobacteria by 16S ribosomal ribonucleic acid sequence comparison: Transfer of Eikenella and Simonsiella to the family Neisseriaceae (emend.). Int. J. Syst. Bacteriol. 39:258–266. Etherington, D. J., P. B. Newman, R. H. Dainty, S. M. Partridge. 1976. Purification and properties of the extracellular metallo-proteinases of Chromobacterium lividum (NCIB 10926). Biochim. Biophys. Acta 445:739–752. Falsen, E. 1989. Catalogue of strains, Culture Collection. University of Göteborg. Göteborg, Sweden. Farrar, W. E., N. M. O’Dell. 1976. β-Lactamase activity in Chromobacterium violaceum. J. Infect. Dis. 134:290– 293. Feldman, R. B., G. A. Stem, C. I. Hood. 1984. Chromobacterium violaceum infection of the eye. Arch. Ophthalmol. 102:711–713. Fujisawa, H., H. Nakata. 1987. Phenylalanine 4-monooxygenase from Chromobacterium violaceum. Methods Enzymol. 142:44–49. Gauthier, M. J. 1976. Morphological, physiological and biochemical characteristics of some violet-pigmented bacteria isolated from seawater. Can. J. Microbiol. 22: 138–149. Gauthier, M. J. 1982. Validation of the name Alteromonas luteoviolacea. Int. J. Syst. Bacteriol. 32:82–86. Gogolewski, R. P. 1983. Chromobacterium violaceum septicaemia in a dog. Aust. Vet. J. 60:226. Gonzalez, C., C. Gutierrez, T. Grande. 1987. Bacterial flora in bottled uncarbonated mineral drinking water. Can. J. Microbiol. 33:1120–1125. Goor, M., E. Falsen, B. Pot, M. Gillis, K. Kersters, J. De Ley. 1986. Taxonomic position of the phytopathogen Pseudomonas rubrisubalbicans and related clinical isolates. 37.Abstr. XIV International Congress of Microbiology, 7–13 September, Manchester, UK. Groscop, J. A., M. N. Brent. 1964. The effects of selected strains of pigmented microorganisms on small freeliving amoebae. Can. J. Microbiol. 10:579–584. Groves, M. G., J. M. Strauss, J. Abbas, C. E. Davis. 1969. Natural infections of gibbons with a bacterium producing a violet pigment (Chromobacterium violaceum). J. Inf. Diseases 120:605–610. Hamada, M., S. Kondo, H. Nakamura, T. Ikeda, D. Ikeda, K. Iinuma, S. Gomi, Y. Ikeda, T. Takeuchi, H. Umezawa, Y. Iitaka. 1983. A new antibiotic, 3,6-dihydroxyindoxazene. J. Antibiot. 36:445–447. Hoshino, T., T. Kondo, T. Uchiyama, N. Ogasawara. 1987. Biosynthesis of violacein a novel rearrangement in tryp-

CHAPTER 3.2.8 tophan metabolism with a 1 2-shift of the indole ring. Agric. Biol. Chem. 51:965–968. Hussain, A., V. Van cˇ ura. 1970. Formation of biologically active substances by rhizosphere bacteria and their effect on plant growth. Folia Microbiologica 15:468– 478. Imai, H., K. Suzuki, S. Miyazaki, K. Tanaka, S. Watanabe, M. Iwanami. 1983. A new antibiotic Y-TO678H produced by a Chromobacterium species. J. Antibiot. 36:911–912. Johnson, E. A., R. J. S. Beer. 1971. Violacein, spectrum no. J8/2. H. M. Perkampus, I. Sandeman, and C. J. Timmons (ed.) UV atlas of organic compounds, vol. 5. Butterworth. London. Johnson, W. M., A. F. Disalvo, R. R. Steuer. 1971. Fatal Chromobacterium violaceum septicemia. Am. J. Clin. Pathol. 56:400– 406. Kaufman, S. C., D. Ceraso, A. Schugurensky. 1986. First case report from Argentina of fatal septicemia caused by Chromobacterium violaceum. J. Clin. Microbiol. 23:956– 958. Keeble, J. R., T. Cross. 1977. An improved medium for the enumeration of Chromobacterium in soil and water. J. Appl. Bacteriol. 43:325–327. Koburger, J. A., S. O. May. 1982. Isolation of Chromobacterium spp. from foods, soil, and water. Appl. Environm. Microbiol. 44:1463–1465. Kwok, O. C. H., P. C. Fahy, H. A. J. Hoitink, G. A. Kuter. 1987. Interactions between bacteria and Trichoderma hamatum in suppression of rhizoctonia damping-off in bark compost media. Phytopathology 77:1206–1212. Laws, L., W. T. R. Hall. 1964. Chromobacterium violaceum infections in a pig. Queensland J. Agric. Sci. 20:393– 400. Lersten, N. R., H. T. Horner, Jr. 1976. Bacterial leaf nodule symbiosis in angiosperms with emphasis on Rubiaceae and Myrsinaceae. Bot. Rev. 42:145–214. Lin, Y. C., A. S. Kester. 1988. Production, location and binding of violacein in Janthinobacterium. Abstr. Annual Meeting of the American Society for Microbiology, Miami Beach, FL. 88:186. Liu, C. H., C. N. Weng, Y. L. Lin, R. M. Chu. 1988. Pathology of experimental infection with Chromobacterium violaceum in pigs. J. Clin. Soc. Vet. Sci. 14:191–202. Logan, N. A. 1989. Numerical taxonomy of violet-pigmented, Gram-negative bacteria and description of Iodobacter fluviatile gen. nov., com. nov. Int. J. Syst. Bacteriol. 39:450– 456. Machin Villafranca, C., M. Ley, L. Torres Hernandez. 1986. Infeccion por Chromobacterium violaceum. Rev. Cubana Med. Trop. 38:353–357. Martin, J. P., S. J. Richards. 1963. Decomposition and binding action of polysaccharide from Chromobacterium violaceum in soil. J. Bacteriol. 85:1288–1294. McClure, H. M., J. Chang. 1976. Chromobacterium violaceum infection in a nonhuman primate (Macaca assamensis). Lab. Anim. Sci. 26:807–810. Miller, D. P., W. T. Blevins, D. B. Steele, M. D. Stowers. 1988. A comparative study of virulent and avirulent strains of Chromobacterium violaceum. Can. J. Microbiol. 34:249– 255. Moffet, M. L., R. R. Colwell. 1968. Adansonian analysis of the Rhizobiaceae. J. Gen. Microbiol. 51:245–266. Moss, M. O., T. N. Bryant. 1982. DNA:rRNA hybridization studies of Chromobacterium fluviatile. J. Gen. Microbiol. 128:829–834. Moss, M. O., C. Ryall. 1981. The genus Chromobacterium. 1355–1364.M. P. Starr, H. Stolps, H. G. Trü per, A. B. Balows, and H. G. Schlegel (ed.) The prokaryotes: A

CHAPTER 3.2.8

The Genera Chromobacterium and Janthinobacterium

handbook in habitats, isolation, and identification of bacteria. Springer-Veriag. Berlin. Moss, M. O., C. Ryall, N. A. Logan. 1978. The classification and characterization of chromobacteria from a lowland river. J. Gen. Microbiol. 105:11–21. Mudarris, M., B. Austin. 1988. Quantitative and qualitative studies of the bacterial microflora of turbot Scophthalmus maximus L. gills. J. Fish. Biol. 32:223–229. Naito, S., I. Shiga, N. Yamaguchi. 1986a. Isolation and identification of violet pigment producing bacteria from yudemen boiled japanese noodle and protection against microbiological deterioration. Part X. J. Jap. Soc. Food Sci. Technol. 33:752–758. Naito, S., I. Shiga, N. Yamaguchi. 1986b. Antimicrobial activity of violet pigment produced by Janthinobacterium lividum isolated from yudemen boiled japanese noodle. Part XIV. J. Jap. Soc. Food Sci. Technol. 33:759–763. Niven, D. F., P. A. Collins, C. J. Knowles. 1975. The respiratory system of Chromobacterium violaceum grown under conditions of high and low cyanide evolution. J. Gen. Microbiol. 90:271–285. Ognibene, A. J., E. Thomas. 1970. Fatal infection due to Chromobacterium violaceum in Vietnam. Am. J. Clin. Pathol. 54:607–610. Onile, A., B. O. Sobowale, T. Odugbemi. 1984. Human infection due to Chromobacterium violaceum: a report from Ilorin, Nigeria. East Afr. Med. J. 61:849–852. Parker, W. L., M. L. Rathnum, J. H. Johnson, J. S. Wells, P. A. Principe, R. B. Sykes. 1988. Aerocyanidin, a new antibiotic produced by Chromobacterium violaceum. J. Antibiot. 41:454– 460. Pember, S. O., J. J. Villafranca, S. J. Benkovic. 1987a. Chromobacterium violaceum phenylalanine 4-monooxygenase. Methods Enzymol. 142:50–56. Pember, S. O., S. J. Benkovic, J. J. Villafranca, M. Passenkiewicz-Gierula, W. E. Antholine. 1987b. Adduct formation between the cupric site of phenylalanine hydroxylase from Chromobacterium violaceum and 6,7-dimethyltetrahydropterin. Biochemistry 26:4477– 4483. Petrillo, V. F., V. Severo, M. M. Santos, E. L. Edelweiss. 1984. Recurrent infection with Chromobacterium violaceum: first case report from South America. J. Infect. 9:167– 169. Quevedo-Sarmiento, J., A. Ramos-Cormenzana, J. GonzalezLopez. 1986. Isolation and characterization of aerobic heterotrophic bacteria from natural spring waters in the Lanjaron area (Spain). J. Appl. Bacteriol. 61:365–372. Rippey, S. R., V. J. Cabelli. 1979. Membrane filter procedure for enumeration of Aeromonas hydrophila in fresh waters. Appl. Environm. Microbiol. 38:108–113. Riveros, R., M. Haun, V. Campos, N. Duran. 1988. Bacterial chemistry IV. Arq. Biol. Techol. (Curitiba) 31:475– 487. Rossau, R., G. Vandenbussche, S. Thielemans, P. Segers, H. Grosch, E. Göthe, W. Mannheim, J. De Ley. 1989. Ribosomal ribonucleic nucleic acid cistron similarities and deoxyribonucleic acid homologies of Neisseria, Kingella, Eikenella, Simonsiella, Alysiella, and Centers for Disease Control groups EF-4 and M-5 in the emended family Neisseriaceae. Int. J. Syst. Bacteriol. 39:185–198. Ryall, C., M. O. Moss. 1975. Selective media for the enumeration of Chromobacterium ssp. in soil and water. J. Appl. Bacteriol. 38:53–59. Sebek, O. K. 1965. Microbiological method for the determination of L-tryptophan. J. Bacteriol. 90:1026–1031.

745

Shetty, M., A. Venkatesh, S. Shenoy, P. G. Shivananda. 1987. Chromobacterium violaceum meningitis: a case report. Indian J. Med. Sci. 41:275–276. Simo, F., P. D. Reuman, F. J. Martinez, E. M. Ayoub. 1984. Chromobacterium violaceum as a cause of periorbital cellulitis. Pediatr. Infect. Dis. 3:561–563. Singh, B. N. 1942. Toxic effects of certain bacterial metabolic products on soil protozoa. Nature (London) 149:168. Singh, B. N. 1945. The selection of bacterial food by soil amoebae, and the toxic effects of bacterial pigments and other products on soil protozoa. Br. J. Exp. Pathol. 26:316–325. Singh, P. D., W.-C. Liu, J. Z. Gougoutas, M. F. Malley, M. A. Porubcan, W. H. Trejo, J. S. Wells, R. B. Sykes. 1988. Aerocavin, a new antibiotic produced by Chromobacterium violaceum. J. Antibiot. 41:446–453. Sivendra, R., S. H. Tan. 1977. Pathogenicity of nonpigmented cultures of Chromobacterium violaceum. J. Clin. Microbiol. 5:514–516. Smith, A. D., R. J. Hunt. 1985. Solubilization of gold by Chromobacterium violaceum. J. Chem. Technol. Biotechnol. 35:110–116. Sneath, P. H. A. 1956. Cultural and biochemical characteristics of the genus Chromobacterium. J. Gen. Microbiol. 15:70–98. Sneath, P. H. A. 1960. A study of the bacterial genus Chromobacterium. Iowa State J. Sci. 34:243–500. Sneath, P. H. A. 1974. Chromobacterium Bergonzini 1881. 354–357.R. E. Buchanan, and N. E. Gibbons (ed.) Bergey’s manual of determinative bacteriology. Williams and Wilkins. Baltimore. Sneath, P. H. A. 1984. Chromobacterium Bergonzini 1881. 580–582.N. R. Krieg, and J. G. Holt (ed.) Bergey’s manual of systematic bacteriology, vol. 1. Williams and Wilkins. Baltimore. Sneath, P. H. A., J. P. F. Whelan, R. B. Singh, D. Edwards. 1953. Fatal infection by Chromobacterium violaceum. Lancet ii:276–277. Sorensen, R. U., M. R. Jacobs, S. B. Shurin. 1985. Chromobacterium violaceum adenitis acquired in the northern United States as a complication of chronic granulomatous disease. Pediatr. Infect. Dis. 4:701–702. Stackebrandt, E., R. G. E. Murray, H. G. Trü per. 1988. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “Purple bacteria and their relatives.” Int. J. Syst. Bacteriol. 38:321–325. Stevens, C. L., P. Blumbergs, F. A. Daniher, R. W. Wheat, A. Kujomoto, E. L. Rollins. 1963. The identification and synthesis of the 4-amino sugar from Chromobacterium violaceum. J. Am. Chem. Soc. 85:3061. Suarez, A. E., B. Wenokur, J. M. Johnson, L. D. Saravolatz. 1986. Nonfatal chromobacterial sepsis. South. Med. J. 79:1146–1148. Umezawa, H., T. Aoyagi, S. Ohuchi, A. Okuyama, H. Suda, T. Takita, M. Hamada, T. Takeuchi. 1983. Arphamenines A and B, new inhibitors of aminopeptidase B, produced by bacteria. J. Antibiot. 36:1572–1575. Van Landschoot, A., J. De Ley. 1983. Intra- and intergeneric similarities of the rRNA cistrons of Alteromonas, Marinomonas (gen. nov.) and some other Gram-negative bacteria. J. Gen. Microbiol. 129:3057–3074. Victoria, B., H. Baer, E. M. Ayoub. 1974. Successful treatment of systemic Chromobacterium violaceum infection. J. Am. Med. Assoc. 230:578–580. Weiss, B., K. S. Hui, M. Hui, A. Lajtha. 1983. Effect of bestatin analogues and other compounds on enkephalin

746

M. Gillis and J. de Ley

hydrolysis by an aminopeptidase from the mesophiles Pseudomonas sp. ATCC 11299A and Chromobacterium violaceum ATCC 12540. Res. Commun. Chem. Pathol. Pharmacol. 39:463– 475. Wilkey, I. S., A. McDonald. 1983. A probable case of Chromobacterium violaceum infection in Australia. Med. J. Aust. 2:39–40. Williams, G. L., L. J. Albright. 1984. The effects of the insecticide acephate on the growth and nutrient uptake of an aquatic bacterium. Can. J. Microbiol. 30:375–380. Woese, C. R., W. G. Weisburg, B. J. Paster, C. M. Hahn, R. S. Tanner, N. R. Krieg, H.-P. Koops, H. Harms, E. Stacke-

CHAPTER 3.2.8 brandt. 1984. The phylogeny of purple bacteria: the beta subdivision. Syst. Appl. Microbiol. 5:327–336. Wu, S. H., S. J. Lin, H. M. Tso, C. B. Liu, W. C. Tsai. 1986. (Fatal septicemia due to Chromobacterium violaceum). Chung Hua Min Kuo Wei Sheng Wu Chi Mien I Hsueh Tsa Chic 19:289–294. Zentmyer, G. A. 1965. Bacterial stimulation of sporangium production in Phytophthora cinnamomi. Science 150:1178–1179. Zwang, J., Z. G. Mai, Z. H. Dung. 1987. Mobilization of granulocyte-macrophage colony forming cell CFU. Acta Physiol. Sin. 39:61–67.

Prokaryotes (2006) 5:747–750 DOI: 10.1007/0-387-30745-1_33

CHAPTER 3.2.9 ehT

areneG

mu i re t cabo l l yhP

dna

mur t caborhcO

The Genera Phyllobacterium and Ochrobactrum JEAN SWINGS, BART LAMBERT, KAREL KERSTERS AND BARRY HOLMES

The study of leaf nodulation in several tropical plant families dates back to the beginning of the 20th century. Various genera of bacteria, e.g., Phyllobacterium, were thought to be associated with leaf nodules (see below). Phyllobacterium was described by Knösel (1962) but the genus only found wide taxonomic recognition with the publication of Bergey’s Manual of Systematic Bacteriology (Knösel, 1984). DNArRNA hybridizations revealed a close relationship between Phyllobacterium and bacteria from the Centers for Disease Control (CDC) group Vd within the “alpha” subclass (rRNA superfamily IV) of the proteobacteria together with those throughout of the genera Brucella, Rhizobium, Mycoplana, and Agrobacterium (De Ley et al., 1987; Stackebrandt et al., 1988). Holmes et al. 1988 proposed the name Ochrobactrum anthropi for CDC group Vd. They also demonstrated that Phyllobacterium showed closest phenotypic similarity to Ochrobactrum anthropi. In the present chapter, the genera Phyllobacterium and Ochrobactrum are treated together because of their close phylogenetic relatedness. Nevertheless, the ecological niches from which these bacteria have been isolated are quite different: Phyllobacterium strains occur in leaf nodules and in the rhizosphere, whereas Ochrobactrum strains have been isolated almost exclusively from human clinical material. In clinical microbiology, Ochrobactrum, more recently described as Achromobacter group Vd until formally named, was usually treated together with Achromobacter xylosoxidans (more recently known as Alcaligenes xylosoxidans subsp. xylosoxidans) (Gilardi, 1978; Rubin et al., 1985). It has now become apparent that Ochrobactrum and Phyllobacterium are not related phylogenetically to Achromobacter xylosoxidans, which belongs to the Alcaligenaceae in the “beta” subclass (rRNA superfamily III) of the proteobacteria (De Ley et al., 1986).

Habitats The Occurrence of Phyllobacterium in Leaf Nodules Plants of over 400 species of the families Rubiaceae and Myrsinaceae are reported to produce leaf nodules. Short-lived bacterial colonies become established intercellularly and die before full leaf expansion (Lersten and Horner,

This chapter was taken unchanged from the second edition.

1976). Bacterial leaf nodules were defined by Horner and Lersten (1972) as: Internal cavities in the leaf lamina, open to the exterior by way of stomatal pores only in early stages of nodule development. The bacteria inhabiting such nodules are part of a population maintained by the host plant in the vegetative and floral buds and passed to the succeeding generations through the seeds. This definition rules out all casual bacteriaplant associations. Despite more than a century of research on leaf nodulation (for a review, see Lersten and Horner, 1976) it is not yet clear which bacteria are involved and how they participate in the nodulation process. In the past, the bacterial symbionts in the nodules were recognized as belonging to the genera Chromobacterium, Xanthomonas, Klebsiella and Phyllobacterium (Horner and Lersten, 1972). Knösel (1962) isolated and described Phyllobacterium and stated that it is able to induce characteristic nodules on leaves (Knösel, 1984), basing his conclusions on earlier investigations by Miehe (1911), von Faber (1912), and de Jongh (1938). However, it is not clear whether the bacteria previously studied by these workers are identical to those described by Knösel. There is no proof for the leaf nodulation capacity of Phyllobacterium.

The Occurrence of Phyllobacterium in the Rhizosphere Pseudomonas fluorescens and Phyllobacterium were found to be the two most frequently occurring bacteria on root surfaces during a large-scale assessment of the rhizobacterial communities of young sugar beet plants (between the second and 10th leaf stage) (Lambert et al., 1990). These bacteria were found in 198 out of 1,100 plants investigated in Belgium and Spain at densities up to 2 × 108 per gram of root. This was the first record of the occurrence of Phyllobacterium in the rhizosphere. Extensive analyses of the microflora from the rhizoplane of other crop plants have not revealed the presence of Phyllobacterium.

748

J. Swings et al.

Occurrence of Ochrobactrum (CDC group Vd) in Clinical Samples and the Environment Ochrobactrum anthropi strains have been isolated predominantly from human blood, urogenital tracts, urine, respiratory tracts, ears, and wounds (Tatum et al., 1974; Holmes et al., 1988). Only a few strains were isolated from feces, spinal fluid, eye, hospital apparatus, or the environment (arsenical cattle-dipping fluid, soil, sewage, and from thin-layer Sephadex plates) (Holmes et al., 1988). The clinical significance of O. anthropi remains largely unknown; strains of this species are considered to be opportunistic pathogens. One strain has been reported in association with a pancreatic abscess (Appelbaum and Campbell, 1980) and another has been associated with a puncture wound leading to osteochondritis of a foot (Barson et al., 1987). Isolation Isolation of Phyllobacterium from the Sugar Beet Rhizosphere (Lambert et al., 1990) The entire root system of a plant is carefully washed to remove adhering soil, then vigorously shaken for 15 min using a flask shaker in a phosphate buffered saline solution containing 0.025% Tween 20. Serial dilutions of the resulting suspensions are plated on 10% trypticase soy broth plus 2% agar (TSBA). After incubation at 28°C for 2 days, Phyllobacterium colonies are beige-colored, large (±5 mm), and convex with an entire edge. They are very mucoid and glistening. Several subcultures are necessary in order to obtain pure isolates. Phyllobacterium readily grows on other nonselective media for aerobic, heterotrophic bacteria (nutrient agar, trypticase soy agar, Luria-Bertani agar, etc.).

Isolation of Phyllobacterium from Leaf Nodules (Knösel, 1984) Washed leaf pieces containing nodules are macerated by rubbing and placed in saline. After shaking, serial dilutions of the suspension are plated onto carrot juice agar containing yeast extract. After incubation at 28°C, a variety of pigmented and nonpigmented colonies develops. Typical colonies are transferred into liquid carrot juice medium which is then used to inoculate the tests used for identification. The cultures should be observed after 24 to 48 h to confirm the presence of star clusters.

Isolation of Ochrobactrum anthropi from Clinical Material Samples from human blood, urine and urogenital tract, sputum, wound, throat, stool, rectum, and other material are plated on blood agar or on MacConkey agar (Rubin et al., 1985; Holmes et al., 1988). Two colony types were recognized on blood agar at 35°C after 48 h: 1) an extremely mucoid type, 0.1 to 4 mm in diameter, opaque and gray-white, showing beta-hemolysis; and 2) a pinpoint type, opaque and semiglossy, gray or white, showing beta-hemolysis after prolonged incubation (Chester and Cooper, 1979). On MacConkey agar after continued incubation, colonies from 3 to 5 mm diameter were formed, opaque and pink to purple. Half the strains produced

CHAPTER 3.2.9 colonies with such notably gummy consistency that it was difficult to remove them from the agar surface (Chester and Cooper, 1979). On MacConkey agar, Ochrobactrum can be easily overlooked due to the small size of the colonies after overnight incubation (Chester and Cooper, 1979).

Preservation of Cultures Well-grown cultures may be kept at 4°C for 2 to 3 months on nutrient agar slants in screw-capped bottles. Isolates can be stored freeze-dried or as a suspension in 25% glycerol at −70°C.

Identification Phyllobacterium and Ochrobactrum are both Gram-negative, strictly aerobic bacteria with a respiratory type of metabolism. They show oxidase and catalase activity and both grow on nutrient agar. Indole is not produced. They oxidize glucose and xylose but not lactose (Clark et al., 1984; G. L. Gilardi, personal communication, Lambert et al., 1990). They do not produce extracellular enzymes for the hydrolysis of Tween 80, DNA, gelatin, starch, or casein (Gilardi, 1978). The API 20NE system will identify them as Achromobacter sp., which is still a “dumping ground” (Kersters and De Ley, 1984) for various aerobic peritrichously flagellated rods. Recent work on Phyllobacterium and Ochrobactrum has led to the differentiation scheme given in Table 1. An extensive description of Phyllobacterium and Ochrobactrum can be found in Lambert et al. (1990) and Holmes et al. (1988), respectively. A comment should be made on the flagellar arrangement of strains of both genera. Phyllobacterium and Ochrobactrum strains both show 1 to 3 polar, subpolar, or Table 1. Features that differentiate Phyllobacterium and Ochrobactrum (CDC group Vd). Property Utilizationa of: L-Citrulline Glutarate Erythritol Glycine L-Tryptophan Hydrolysisb of paranitrophenol-αmaltoside Hydrolysisb of paranitrophenol-αxylopyranoside

Phyllobacteriumc

Ochrobactrumd

− − − − + +

+ + + + − −

+

−

Using API 50CH and 50AO, 50AA strips (API System, La Balme-les-Grottes, Montalieu-Vercieu, France). Using API ZYM strips. 15 strains total. From Lambert et al. (1990). 56 strains total. From Holmes et al. (1988).

CHAPTER 3.2.9

The Genera Phyllobacterium and Ochrobactrum

lateral flagella (Lambert et al., 1990). According to Holmes et al. 1988 and Gilardi (1978) Ochrobactrum strains are characterized by peritrichous flagella, but other workers have found only polar, subpolar, or lateral flagella but no definitely peritrichous flagella (Clark et al., 1984; Chester and Cooper, 1979). An unambigous identification of Phyllobacterium or Ochrobactrum is possible by application of carbon assimilation tests (API 50CH, 50AO, or 50AA) or various conventional tests (Tatum et al., 1974; Rubin et al., 1985; Holmes et al., 1988; Lambert et al., 1990) or by protein electrophoretic fingerprinting. Variability in Gram reaction, the presence of curved cells with swollen-ends, and the formation of extremely small colonies after overnight incubation, which develop into mucoid colonies upon continued incubation, may lead to incorrect identification as Corynebacterium or Klebsiella (Chester and Cooper, 1979). Knösel (1984) used the criterium of nitrate reduction to differentiate Phyllobacterium myrsinacearum (+) from P. rubiacearum (−). Lambert et al. (1990), however, found that P. rubiacearum LMG 1t1T reduced nitrates. Using the API 2ONE system, they found that all Phyllobacterium strains showed a slight NO3− reduction. These observations cast serious doubt on the usefulness of this criterion for species differentiation within the genus Phyllobacterium. All Ochrobactrum strains are capable of nitrate reduction (Gilardi, 1978; Clark et al., 1984; Holmes et al., 1988).

al., 1990), which occur in all the classical nitrogen-fixing bacteria. Phyllobacterium is able to interact with plant tissues, as demonstrated by the tumor induction by Ti-plasmid-carrying Phyllobacterium strains on Kalanchoe plants (Lambert et al., 1990). Tumor induction involves the attachment of bacterial cell wall sites to plant cell wall sites prior to the induction of T-DNA transfer. The chromosomal genes chvA, chvB, exoC, and att are the only ones found to be involved in attachment but could not be detected in Phyllobacterium strains, suggesting that other genes are present with similar functions. There are no data indicating that Phyllobacterium is pathogenic or deleterious to plants. A striking feature, common to both Phyllobacterium and Ochrobactrum, is nutritional versatility, which makes their rapid growth and proliferation possible in rich environments such as the root surface and clinical sites. A number of Phyllobacterium isolates showed antifungal and antibacterial activities (Lambert et al., 1990). The oxidation of lactose to 3-ketolactose is a unique feature of many Agrobacterium strains (Kersters and De Ley, 1984) and is negative for all Phyllobacterium and Ochrobactrum strains.

Physiological Properties Cellular fatty acids of Ochrobactrum anthropi (Dees and Moss, 1978) as well as antimicrobial susceptibilities have been determined (von Graevenitz, 1985; Gilardi, 1989). Growth of Phyllobacterium strains was inhibited by doxycycline, novobiocin, framycetin, and tetracycline (Lambert et al., 1990). The most intriguing question that remains unanswered is whether Phyllobacterium has a real symbiotic cyclical relationship with its hosts. Reinfection studies did not allow unambiguous conclusions to be made. It has also been suggested that leaf nodule bacteria produce plant growth hormones, particularly cytokinins, which are necessary for the normal functioning of the plant (Fletcher and Rhodes-Roberts, 1976; reference is not an exact matchRodrigues-Pereira et al., 1972). Other authors claimed that leaf nodule bacteria can fix nitrogen, but this was contested by Van Hove (1976) and Lersten and Horner (1976). The lack of ability to fix nitrogen is also indicated by the absence of the nif HDK-like genes (Lambert et

749

Applications No applications of the genera Phyllobacterium or Ochrobactrum are yet known. However, the fact that Phyllobacterium is a predominant bacterium on the root surface of sugar beet plants, its capacity to “communicate” with plant tissues, and its non-pathogenic status make this bacterium an interesting new candidate for use in plant growth promotion or biological control of soil-borne diseases. Indeed, some of the sugar beet isolates exert a broad-spectrum antifungal activity against major phytopathogenic fungi.

Literature Cited Appelbaum, P. C., D. B. Campbell. 1980. Pancreatic abscess associated with Achromobacter group Vd biovar 1. J. Clin. Microbiol. 12:282–283. Barson, W. J., B. A. Cromer, M. J. Marcon. 1987. Puncture wound osteochondritis of the foot caused by CDC group Vd. J. Clin. Microbiol. 25:2014–2016. Chester, B., L. H. Cooper. 1979. Achromobacter species (CDC group Vd): morphological and biochemical characterization. J. Clin. Microbiol. 9:425–436. Clark, W. A., D. G. Hollis, R. E. Weaver, P. Riley. 1984. Identification of unusual pathogenic Gram-negative aerobic and facultatively anaerobic bacteria. Centers for Disease Control, Atlanta.

750

J. Swings et al.

Dees, S. B., C. W. Moss. 1978. Identification of Achromobacter species by cellular fatty acids and by production of keto acids. J. Clin. Microbiol. 8:61–66. De Jongh, P. 1938. On the symbiosis of Ardisia crispa (Thunb.). A. DC. Verh. Kon. Ned. Akad. Wetensch. Afd. Natuurk. Tweede Sect. II 37:1–74. De Ley, J., W. Mannheim, P. Segers, A. Lievens, M. Denijn, M. Vanhoucke, M. Gillis. 1987. Ribosomal ribonucleic acid cistron similarities and taxonomic neighborhood of Brucella and CDC Group Vd. Int. J. Syst. Bacteriol. 37:35–42. De Ley, J., P. Segers, K. Kersters, W. Mannheim, A. Lievens. 1986. Intra- and intergeneric similarities of the Bordetella ribosomal ribonucleic acid cistrons: proposal for a new family, Alcaligenaceae. Int. J. Syst. Bacteriol. 36:405–414. Fletcher, L. M., M. E. Rhodes-Roberts. 1976. The bacterial leaf nodule association in Psychotria. 99–118. D. W. Lovelock (ed.) Soc. appl. bacteriol. Technical series no 12. Academic Press, London. Gilardi, G. L. 1978. Identification of miscellaneous glucose non-fermenting Gram-negative bacteria. 45–65. G. L. Gilardi (ed.) Glucose non-fermenting gram-negative bacteria in clinical microbiology. CRC Press, Inc. Boca Raton, FL. Holmes, B., M. Popoff, M. Kiredjian, K. Kersters. 1988. Ochrobactrum anthropi gen. nov., sp. nov. from human clinical specimens and previously known as group Vd. Int. J. Syst. Bacteriol. 38:406–416. Horner, H. T., N. R. Lersten. 1972. Nomenclature of bacteria in leaf nodules of the families Myrsinaceae and Rubiaceae. Int. J. Syst. Bacteriol. 22:117–122. Kersters, K., J. De Ley. 1984. Genus III. Agrobacterium. 244– 254. N. R. Krieg and J. G. Holt (ed.) Bergey’s manual of systematic bacteriology, vol. 1. Williams & Wilkins, Baltimore. Knösel, D. 1962. Prü fung von Bakterien auf Fähigkeit zur Sternbildung. Zentralbl. Bakteriol. Parasitenk. Infektionskr. Hyg. Abt. 2 116:79–100. Knösel, D. 1984. The genus Phyllobacterium. 254–256. N. R. Krieg, and J. G. Holt (ed.) Bergey’s manual of systematic bacteriology, vol. 1. Williams & Wilkins, Baltimore.

CHAPTER 3.2.9 Lambert, B, H. Joos, S. Dierickx, R. Vantomme, J. Swings, K. Kersters, M. Van Montagu. 1990. The identification and plant interaction of a Phyllobacterium sp. a predominant rhizobacterium of young sugar beet plants. Appl. Environ. Microbiol. 56:1093–1102. Lersten, N. R., H. T. Horner, Jr. 1976. Bacterial leaf nodule symbiosis in angiosperms with emphasis on Rubiaceae and Myrsinaceae. Bot. Rev. 42:145–214. Miehe, H. 1911. Die Bakterienknoten an den Blatträndern der Ardisia crispa A. DC. Javanische Studien V. Abh. Math. Phys. Kl. Königl. Sächs. Gesellsch. Wiss. Leipzig, 32:399–431. Rodriguez-Pereira, A. S., P. J. W. Houwen, H. W. J. Deurenberg-Vos, B. F. Dey. 1972. Cytokinins and the bacterial symbiosis of Ardisia species. Zeitschr. Pflanzenphysiol. 68:170–177. Rubin, S. J., P. A. Granata, B. L. Wasilanskas. 1985. Glucosenonfermenting Gram-negative bacteria. 330–349. A. Balows, W. J. Hausler, Jr., and H. J. Shadomy (ed.) Manual of clinical microbiology, 4th ed. American Society for Microbiology. Washington, D.C. Stackebrandt, E., R. G. E. Murray, H. G. Trü per. 1988. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “purple bacteria and their relatives.” Int. J. Syst. Bacteriol. 38:321–325. Tatum, H. W., W. H. Ewing, R. E. Weaver. 1974. Miscellaneous Gram-negative bacteria. 270–294. E. H. Lennette, E. H. Spaulding, and J. P. Truant (ed.) Manual of clinical microbiology, 2nd ed. American Society for Microbiology. Washington, D.C. Van Hove, C. 1976. Bacterial leaf symbiosis and nitrogen fixation. 551–560. P. S. Nutman (ed.) Symbiotic nitrogen fixation in plants. Cambridge University Press. London, U.K. von Faber, F. C. 1912. Das erbliche Zusammenleben von Bakterien und tropischen Pflanzen. Jahrb. Wiss. Bot. 51:285–375. von Graevenitz, A. 1985. Ecology, clinical significance, and antimicrobial susceptibility of infrequently encountered glucose-nonfermenting Gram-negative rods. 181–232. G. L. Gilardi (ed.) Nonfermentative Gram-negative rods. Laboratory identification and clinical aspects. Marcel Dekker, Inc.. New York.

Prokaryotes (2006) 5:751–757 DOI: 10.1007/0-387-30745-1_34

CHAPTER 3.2.10 ehT

suneG

a i xreD

The Genus Derxia JAN HENDRIK BECKING

In the genus Derxia only a single species is recognized, D. gummosa. This species was originally described by Jensen et al. (1960) from an isolate obtained from Indian soil, but was later found to be widely distributed in soils of South America (Brazil), Southern Africa, Indonesia, and China, but not in temperate regions. Although Roy and Sen (1962) described a second species of Derxia (D. indica sp. nov.) from a sample of partially retted jute plant (Corchorus olitorius L.) from Uttar Pradesh, India, they did not present cultural and physiological data showing that their isolate was sufficiently different from Derxia gummosa to warrant the designation of a new species. The genus is named for H. G. Derx (1894–1953), a Dutch microbiologist. Using rRNA cistron similarity as a criterion for genetic relatedness, De Smedt et al. (1980) found that D. gummosa is quite different from species of the genera Azotobacter, Azomonas, and Beijerinckia. From hybridization studies with DNAs from three D. gummosa strains using the 14C-labeled rRNAs from a great variety of other organisms, it was quite obvious that the Derxia rRNA cistrons most closely resemble those of Pseudomonas acidovorans, P. solanacearum, Chromobacterium violaceum, Janthinobacterium lividum, and Alcaligenes faecalis. These taxa, together with a few others, constitute the third rRNA superfamily of De Ley (see The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition). Members of these taxa have in common that they consist of Gram-negative rods, usually 1.0–4.0 µm in length and 0.5–1.0 µm in diameter, do not possess resting stages, have a GC content of the DNA of 57–72 mol%, are chemoheterotrophic, exhibit respiratory (oxidative) metabolism, and occur in soil and water habitats. The third rRNA superfamily of De Ley (De Smedt et al., 1980) is identical with the beta subclass of the Proteobacteria, according to the phylogenetic taxon nomenclature of Stackebrandt et al. 1988. Based on numerical analysis of a large number of attributes, Thompson and Skerman (1979) showed fusion of their five Derxia strains tested with “Azotomonas” (“Azotomonas insolita” Stapp, 1940, sometimes termed “Pseudomonas insolita” [Stapp] Brisou, 1961) at group 1296 in their dendrograms of hierarchical interrelations (see also Krieg and Holt, 1984). This indicates that Derxia is not related to the genera Azotobacter or Azomonas and thus confirms the conclusions of De Smedt et al. (1980).

Habitats Derxia is primarily a soil organism occurring in tropical soils. There is no plausible explanation This chapter was taken unchanged from the second edition.

for why this organism is exclusively found in tropical environments and is apparently absent from temperate soils. It has not yet been recorded from aquatic habitats, although it is likely that it also occurs there. Such habitats in these regions have, however, not yet been thoroughly investigated using conditions selective for this organism.

Isolation The original isolate was obtained from a West Bengal (Adisaptagram, India) soil of pH 6.5 by the sieved-soil plate method (see “General Enrichment Procedures” in The Family Azotobacteraceae in the second edition) using plates with a nitrogen-free, mineral mannitol (2%) agar, incubated at room temperature (about 25°C). Yellowish colonies appeared after 5 days around some of the soil particles; these colonies gradually increased in size and finally assumed a rust-brown color. Isolation and further purification were performed by repeated transfer and plating on the same nitrogen-free mannitol agar. Campêlo and Döbereiner (1970) obtained this organism from Brazilian soils also with the sieved-soil plate method, utilizing a nitrogenfree, mineral starch (2%) medium. This medium was a modification of a medium used by Lipman (1903), supplemented with NaHCO3 (0.1 or 0.01%). Isolation Medium for Derxia gummosa (Campêlo and Döbereiner, 1970; Modified from Lipman, 1903) Distilled water Starch K2HPO4 KH2PO4 MgSO4 ·7H 2O CaCl2 NaHCO3 FeCl3 (10%, aqueous solution) Na2MoO4 ·2H 2O Bromothymol blue (0.5%, ethanol solution) Agar

1 liter 20.0 g 0.05 g 0.15 g 0.20 g 0.02 g 0.1 g 1 drop 0.002 g 5 ml 20.0 g

In a survey of 100 samples of Brazilian soil from the states of São Paulo, Rio de Janeiro,

752

J.H. Becking

CHAPTER 3.2.10

Pernambuco, and Para, Campêlo and Döbereiner (1970) found that 36% of the soils tested were positive for this organism. Soil humidity seems to favor the presence of Derxia, since 77% of the flooded soils, 36% of the humid soils, and only 13% of the drier soils tested contained this organism. In a comparative study with agar plates seeded with soil or with root pieces of various plants (mostly Gramineae) of the same locality, Derxia appeared more frequently on the plates inoculated with roots (33%) compared to those inoculated with soil (26%). D. gummosa was found to be present in soils ranging in pH from 4.5 to 6.5, but was observed to be most frequent in soils of pH 5.1–5.5. This species was isolated by the author from various soils of Indonesia, China, Brazil, tropical Africa, and South Africa with the sieved-soil plate method using nitrogen-free, mineral, glucose (1–2%) agar (J. H. Becking, unpublished observations). Derxia strains are relatively acid tolerant and were therefore also isolated by the author from cultures designed to enrich for Beijerinckia strains using a liquid, acidic, nitrogen-free, glucose medium. This liquid medium has the following composition: Enrichment Medium for Derxia gummosa (J. H. Becking, unpublished observations) Distilled water Glucose KH2PO4 MgSO4 ·7H 2O

1 liter 20.0 g 1.0 g 0.5 g

Adjust to pH 5.0.

The medium is dispensed into petri dishes so as to form a thin layer 2–3 mm deep in order to maintain aerobic conditions. The liquid medium is inoculated with soil particles and incubation is at 25° or 30°C. In the sieved-soil plate method mentioned for the isolation, soil particles are evenly distributed over the surface of nitrogen-free mineral agar containing mannitol (Jensen et al., 1960), glucose (J. H. Becking, unpublished observations), or starch (Campêlo and Döbereiner, 1970). It should be noted that mannitol is not utilized by all Derxia gummosa strains. Of the six strains tested by Thompson and Skerman (1979), three did not use mannitol. Moreover, some strains, including the type strain, produced only very scant growth on starch. All strains did, however, utilize glucose as sole source of carbon. Therefore for general practice of the sievedsoil plate method, the following glucose mineral agar medium is recommended: Nitrogen-free, Mineral Glucose Agar for the Isolation of Derxia gummosa with the Sieved-Soil Plate Method (g/liter of distilled water):

Glucose K2HPO4 KH2PO4 MgSO4 ·7H 2O FeCl3 ·6H 2O Na2MoO4 ·2H 2O CaCl2 Agar

20.0 0.8 0.2 0.5 0.025 (or 0.05) 0.005 0.05 15.0

Adjust to pH 6.9.

On the agar plates seeded with the soil particles, yellowish colonies develop around the soil particles. These colonies eventually become larger and acquire a rust-brown color. Isolates should be further purified by repeated streaking on new agar plates. For agar plates obtained from liquid enrichment cultures, one should look for slimy, yellowish colonies with a rather plicated surface (see Fig. 5).

Cultivation For routine maintenance, the following nitrogenfree, mineral glucose medium can be used (g/liter of distilled water): Nitrogen-free, Mineral Glucose Agar Medium Glucose K2HPO4 MgSO4 ·7H 2O NaCl FeSO4 ·7H 2O CaCl2 or CaCO3 Na2MoO4 ·2H 2O Agar

10.0 (or 20.0) 0.5 0.25 (or 0.2) 0.25 0.1 0.1 0.005 15.0

Adjust to pH 6.9.

Jensen et al. (1960) used a nitrogen-free, mineral mannitol agar of the following composition (g/liter of distilled water): Nitrogen-free, Mineral Mannitol Agar Medium Mannitol K2HPO4 MgSO4 ·7H 2O CaCl2 CaCO3 Na2WO4 ·2H 2O FeCl3 and Na2MoO4 ·2H 2O Agar

10.0 0.5 0.2 0.1 5.0 0.0005 trace 15.0.

The medium recommended by Campêlo and Döbereiner (1970) contained starch as carbon source (g/liter of distilled water): Nitrogen-free, Mineral Starch Agar Medium Starch K2HPO4 KH2PO4 MgSO4 ·7H 2O CaCl2 NaHCO3 FeCl3 solution (10%, aqueous solution)

20.0 g 0.05 g 0.15 g 0.2 g 0.02 g 0.1 g 1 drop

CHAPTER 3.2.10 Na2MoO4 ·2H 2O Bromthymol blue solution (0.5%, ethanol solution) Agar

The Genus Derxia 0.002 g 5.0 ml 20.0.

As already stated before, not all Derxia gummosa strains can utilize mannitol as sole source of carbon, while with starch as the carbon source, some strains only produce scant growth.

753

of Derxia acquire on aging a typical dark rustbrown or mahogany-brown color. In addition, in contrast to Azotobacter, Azomonas and Beijerinckia, Derxia cells are catalase negative. In contrast to the spiral-shaped or vibrioid cells of Azospirillum (see The Genera Azospir-

Preservation of Cultures The maintenance procedures of Derxia gummosa are the same as those described for the genus Beijerinckia (see The Genus Beijerinckia in Volume 5).

Identification Derxia can be distinguished from other genera of N2-fixing bacteria by its very slimy (gummy) growth, both on agar plates and in liquid media, combined with the very pleomorphic appearance of the cells (Figs. 1 to 2, 3, 4) depending on age and type of medium. On agar media Azotobacter and Azomonas species never produce such slimy colonies as those of Derxia (Fig. 5). Confusion with Beijerinckia is unlikely, since Beijerinckia cells usually show very characteristic cells with only two polar lipoid bodies, whereas Derxia cells contain numerous lipoid bodies. Moreover, in contrast to Beijerinckia colonies, the colonies

Fig. 2. Three-week-old cells of Derxia gummosa on nitrogenfree glucose agar, showing shrinkage of the cells. Phase contrast microscopy. Bar = 10 µm.

Fig. 1. Seven-day-old cells of Derxia gummosa on nitrogenfree agar containing 2% glucose. Phase contrast microscopy. Bar = 10 µm.

Fig. 3. Ten-day-old cells of Derxia gummosa on peptone agar with 2% glucose. Phase contrast microscopy. Bar = 10 µm.

754

J.H. Becking

CHAPTER 3.2.10

a

b

Fig. 5. Colony of Derxia gummosa on nitrogen-free glucose agar with calcium carbonate showing the coarse and wrinkled colony surface. Bar = 1 cm.

Fig. 4a. Three-month-old cells of Derxia gummosa on nitrogen-free glucose agar. The cells show shrinkage and are enclosed by a thick slime envelope. Phase contrast microscopy. Bar = 10 µm. Fig. 4b. More pronounced shrinkage of cells of a four-month-old Derxia gummosa culture on nitrogen-free glucose agar. Note the thick slime envelopes enclosing the rows of cells. Bar = 10 µm.

illum and Herbaspirillum in the second edition), Derxia cells are mainly straight rods and the lipoid bodies are more numerous. Derxia colonies are far more slimy than those of Azospirillum, and the mahogany-brown color they eventually acquire is in contrast to the pink or whitish color of Azospirillum colonies. Moreover, there are some minor physiological differences. Derxia strains can exhibit nitrogendependent growth on glucose, whereas only some strains of Azospirillum (i.e., only those belonging to the species A. lipoferum) can do so. On the other hand, Derxia strains generally do not grow well on malate, whereas all strains of Azospirillum can grow well on this carbon source. Moreover, Derxia strains are catalase negative, whereas Azospirillum strains give reactions that range from strong to undetectable.

Unlike Azotobacter, Azomonas, and Beijerinckia species, Derxia species can grow as a facultative hydrogen autotroph. In this ability, they show affinities to various other facultative hydrogen autotrophs that can also use H2 to provide energy and reducing power for growth and CO2 fixation, i.e., various Pseudomonas and Alcaligenes species (“Hydrogenomonas” species) (Buchanan and Gibbons, 1974), Xanthobacter species (Wiegel et al., 1978; Wiegel and Schlegel, 1984; Malik and Claus, 1979), nitrate-reducing Paracoccus species (Rittenberg, 1969), Bradyrhizobium japonicum (Hanus et al., 1979), and Azospirillum lipoferum (Malik and Schlegel, 1981; Sampaio et al., 1981) (see also The Mesophilic Hydrogen-Oxidizing (Knallgas) Bacteria in the second edition). For further details on facultative hydrogen autotrophy of Derxia, see section “Physiological and Biochemical Properties” in this Chapter.

Species Description Derxia gummosa Jensen, Petersen, De and Bhattacharya, 1960. Cells are Gram-negative, rod-shaped with rounded ends, 3.0–6.0 µm in length, and 1.0–

CHAPTER 3.2.10

1.2 µm wide, occurring singly or in short chains. Cells are rather pleomorphic, depending on age and the medium. In aging cultures, cells often remain together, forming long filaments of sometimes locally swollen or distorted cells. Some cells may assume enormous sizes (up to 30 µm). Young cells have a homogeneous cytoplasm; older cells typically show large refractile bodies throughout the whole cell. Resting stages are not known. Cells are motile by a short polar flagellum; motile cells are numerous in liquid glucose media containing combined nitrogen, but are rare on nitrogen-deficient solid media. The organism is aerobic, having a strictly respiratory type of metabolism with oxygen as the terminal electron acceptor. Molecular nitrogen is fixed under aerobic conditions and also under decreased oxygen pressures (microaerophilic conditions). Optimum temperature is 25–35°C; growth is slow at 15°C, feeble at 40°C; no growth occurs at 50°C. Growth occurs from pH 5.5 to about 9.0; no growth occurs at pH 4.4. Liquid cultures turn into a gelatinous mass, but growth near the surface is more luxuriant and forms a thick, tough pellicle. Colonies on agar media are at first slimy and semitransparent, later massive and opaque, highly raised with a wrinkled surface. Older colonies develop a rust-brown to dark mahoganybrown color. They are catalase negative. A wide range of sugars, alcohols, and organic acids are oxidized, mostly to CO2, but a small amount of acid, probably acetic acid, is produced during growth in an alkaline medium. They can grow as a facultative hydrogen autotroph. The GC content of the DNA in three strains of Derxia gummosa ranges from 69.2–72.6 mol% (Tm) (De Smedt et al., 1980). In the type strain, it is 70.4 ± 1.7 mol% (Tm) (De Ley and Park, 1966). The appearance of cells from young cultures is depicted in Fig. 1. Older cells on sugar-rich media contain large refractile bodies. Especially on glucose-peptone agar, very elongated cells are produced containing many refractile bodies (Fig. 3). The refractile material is probably poly-βhydroxybutyrate, since it stains with Sudan III and Sudan black, but some vacuoles which do not stain may also be involved. On nitrogen-free glucose agar, older cells undergo shrinkage (Fig. 2) and are finally enclosed by a slime envelope (Fig. 4). Motile cells may become numerous in liquid glucose media containing ammonia or glutamate as the nitrogen source. The cells usually have a single polar flagellum, but some may have a flagellum at each pole. According to Thompson and Skerman (1979), the single polar flagellum is less than a full wave and is less than 3 µm in length. Growth in liquid media usually starts as a ring at the glass-liquid interface and develops into a thick, wrinkled, tough pellicle. Shallow layers of

The Genus Derxia

755

medium change into a firm gelatinous mass after a couple of weeks. The color gradually becomes a dark red-brown. Growth on nitrogen-deficient agar media begins as thin, whitish, or semitransparent scattered colonies. Later, more massive, highly raised, or dome-shaped colonies emerge which rapidly assume a diameter of 1 cm or more (giant colonies) (Fig. 5). These colonies are very reminiscent of those of Beijerinckia species. Colonies are at first whitish or dull yellow with a smooth surface, but the surface soon becomes coarse and wrinkled, and the color deepens to a dark mahogany brown. The slime of these colonies is very tenacious and gumlike, but in the other developmental stages, it is softer and smeary.

Physiological and Biochemical Properties In the original description, Jensen et al. 1960 mentioned the appearance of Derxia gummosa on nitrogen-free, mineral glucose agar as a few “massive” colonies among many “thin,” whitish colonies. The type of colony used as inoculum affected neither the relative numbers nor the distribution of the two colony types. They reported further that the “massive” colonies fixed nitrogen, whereas the “thin” colonies did not. Tchan and Jensen (1963) observed that the formation of “massive” colonies was stimulated by the addition of small quantities of combined nitrogen to the medium and that dinitrogen fixation could not be initiated in a medium completely free of combined nitrogen. Hill and Postgate (1969) found that, with a small quantity of combined nitrogen, batch cultures of Derxia gummosa in otherwise nitrogenfree media could be established with agitation in air. However, this supplement was unnecessary if the cultures were left stagnant or if the atmospheric oxygen concentration was lowered to 0.1 atm or less. Once established, continuous cultures could be maintained in air, provided vigorous stirring was avoided. Hill (1971) demonstrated in more detail that D. gummosa is very sensitive to oxygen and that the presence of oxygen markedly affects its colony morphology. At an oxygen concentration of less than 0.2 atm, only the “massive”-colony type is produced on nitrogen-free, mineral agar media; the “thin”colony type which predominates in air was absent. Only the “massive” colonies reduced acetylene. Apparently, the copious production of a viscous and tenacious slime inhibits the penetration of oxygen and therefore stimulates nitrogen fixation. The colonial dimorphism on agar probably arises because only when the local oxygen concentrations on the agar surface are

756

J.H. Becking

depressed can dinitrogen fixation and subsequent growth to the “massive” colony type take place. The efficiency of nitrogen fixation by D. gummosa varies between 9 and 25 mg N/g of glucose consumed, but in most strains it is distinctly lower than in Azotobacter or Beijerinckia. Growth on nitrogen-deficient media or under nitrogen-fixing conditions is stimulated by small amounts of combined nitrogen (nitrate, ammonia), particularly at the start (Tchan and Jensen, 1963). There is no requirement for amino acids under nitrogen-fixing conditions. However, small amounts of yeast extract are stimulatory for growth under nitrogen-fixing conditions (J. H. Becking, unpublished observations), and although there is no apparent requirement for vitamins or growth factors, it is possible that small amounts of biotin are stimulatory for growth of some strains (J. H. Becking, unpublished observations). As sole source of carbon, growth is good to excellent on glucose, fructose, ethanol, glycerol, mannitol (only utilized by some strains), and sorbitol; growth on lactate is scant. No growth or only a trace of growth occurs on lactose, galactose, maltose, sucrose, formate, acetate, propionate, pyruvate, succinate, malate, fumarate, dulcitol, and sometimes starch. Butyrate, citrate, benzoate, and xylose suppress growth. Growth on methane or methanol as the sole carbon source has also been demonstrated for D. gummosa (Sampaio et al., 1981). Growth with combined nitrogen sources is much faster than with molecular nitrogen and is completely uniform, in contrast to the uneven growth on nitrogen-free agar. Colonies change from a pale yellow through rust-brown to almost black (darkest if nitrate is present), and sometimes a light brown, water-soluble pigment is produced. Growth with glutamic acid, ammonium acetate, alanine, sodium nitrate, and urea decreases from abundant to good in approximately the same sequence. Aspartic acid, asparagine, and peptone give a much slower growth, which is uneven and mostly confined to scattered colonies. Glycine seems to be toxic. Nitrate is not reduced to nitrite or N2 in a glucose-nitrate medium. Indole is not produced from tryptophan. Trace elements, particularly molybdenum, are required for dinitrogen fixation, but vanadium cannot replace molybdenum (Jensen et al., 1960). Thus, Derxia apparently does not have an alternative vanadium-activated nitrogenase. D. gummosa has been shown to grow as a hydrogen autotroph in an atmosphere containing H2 + CO2 + O2, with either N2 or NH4+ as the nitrogen source (Pedrosa et al., 1980). Indeed, it appears to grow nearly as well autotrophically as

CHAPTER 3.2.10

it does heterotrophically. Ribulose-1,5-bisphosphate carboxylase activity, which mediates CO2 fixation, occurs in autotrophically-grown cells but not in cells grown heterotrophically. A medium for testing the autotrophy has the following composition (g/liter of distilled water): Medium for Testing Autotrophy KH2PO4 K2HPO4 MgSO4 ·7H 2O NaCl CaCl2 ·2H 2O FeSO4 ·7H 2O Trace element solution (see below) Biotin Agar

1.2 0.8 0.2 0.2 0.02 0.002 2.0 ml 10 µg 15.0

Trace element solution (g/liter) Na2MoO4 ·2H 2O MnSO4 ·H 2O H3BO3 CuSO4 ·5H 2O ZnSO4 ·7H 2O

1.0 1.75 1.4 0.04 0.12

This medium was employed by Pedrosa et al. (1980) in their studies on the autotrophy of Derxia. In these studies, a very low concentration of potassium malate (0.1 g/liter) was sometimes added to the medium. It should be noted that most strains of Derxia (including the type strain) give only scant growth with malate. Thompson and Skerman (1979) reported no growth on DLmalate in all of the six strains that they tested.

Applications No studies have yet been done on the occurrence and significance of this organism in the rhizosphere of crop plants. However, Campêlo and Döbereiner (1970) found Derxia more frequently on agar plates inoculated with roots (33% positive for Derxia) compared to those inoculated with soil (26%).

Literature Cited Brisou, B. 1961. Etudes de quelques Pseudomonas chromogènes isolés àDiego-Suarez. Bull. Soc. Path. Exot. 54:746–755. Buchanan, R. E., N. E. Gibbons (ed.). 1974. Bergey’s manual of determinative bacteriology. 8th ed. Williams and Wilkins. Baltimore. Campêlo, A. B., Döbereiner, J. A. 1970. Ocorrência de Derxia sp. em solos de alguns Estados Brasileiros. Pesquisa Agropecuá ria Brasileira 5:327–332. De Ley, J., Park, I. W. 1966. Molecular biological taxonomy of some free-living nitrogen-fixing bacteria. Antonie van Leeuwenhoek, Journal of Microbiology and Serology 32:6–16. De Smedt, J., Bauwens, M., Tytgat, R., De Ley, J. 1980. Intraand intergeneric similarities of ribosomal ribonucleic

CHAPTER 3.2.10 acid cistrons of free-living, nitrogen-fixing bacteria. Int. J. Syst. Bacteriol. 30:106–122. Hanus, F. J., Maier, R. J., Evans, H. J. 1979. Autotrophic growth of H2-uptake-positive strains of Rhizobium japonicum in an atmosphere supplied with hydrogen gas. Proc. Natn. Acad. Sci. USA. 76:1788–1792. Hill, S. 1971. Influence of oxygen concentration on the colony type of Derxia gummosa grown on nitrogen-free media. J. Gen. Microbiol. 67:77–83. Hill, S., Postgate, J. R. 1969. Failure of putative nitrogenfixing bacteria to fix nitrogen. J. Gen. Microbiol. 58:277– 285. Jensen, H. L., Petersen, E. J., De, P. K., Bhattacharya, R. 1960. A new nitrogen-fixing bacterium: Derxia gummosa nov. gen. nov. spec. Arch. Microbiol. 36:182– 195. Krieg, N. R., Holt, J. G. (ed.). 1984. Bergey’s manual of systematic bacteriology, vol. 1. Williams and Wilkins. Baltimore. Lipman, J. G. 1903. Experiments on the transformation and fixation of nitrogen by bacteria. 217–285. 16th Annual Report over 1903 of the New Jersey State Agricultural Experiment Station. USA. Malik, K. A., Claus, D. 1979. Xanthobacter flavus, a new species of nitrogen-fixing hydrogen bacteria. Int. J. Syst. Bacteriol. 29:283–287. Malik, K. A., Schlegel, H. G. 1981. Chemolithotropic growth of bacteria able to grow under N2-fixing conditions. FEMS Microbiol. Lett. 11:63–67. Pedrosa, F. O., Döbereiner, J., Yates, M. G. 1980. Hydrogendependent growth and autotrophic carbon dioxide fixation in Derxia. J. Gen. Microbiol. 119:547–551.

The Genus Derxia

757

Rittenberg, S. C. 1969. The roles of exogenous organic matter in the physiology of chemolithotrophic bacteria. Adv. Microbiol. Physiol. 3:159–196. Roy, A. B., Sen, S. 1962. A new species of Derxia. Nature London, 194:604–605. Sampaio, M.-J. A. M., da Silva, E. M. R., Döbereiner, J., Yates, M. G., Pedrosa, F. O. 1981. Autotrophy and methylotrophy in Derxia gummosa, Azospirillum brasilense and A. lipoferum. 447. Gibson, A. H., and Newton, W. E. (ed.) Current perspectives in nitrogen fixation, Proc. 4th Int. Symp. on Nitrogen Fixation, Canberra, Australia, Dec. 1–5, 1980. Elsevier/North-Holland Biomedical Press. Amsterdam. Stackebrandt, E., Murray, R. G. E., Trü per, H. G. 1988. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “purple bacteria and their relatives.” Int. J. Syst. Bacteriol. 38:321–325. Stapp, C. 1940. Azotomonas insolita, ein neuer aerober stickstoffbindender Mikroorganismus. Zentralblatt fü r Bakteriologie, Parasitenkunde, Infektionskrankheiten, und Hygiene, Abt. II 102:142–150. Tchan, Y. T., Jensen, H. L. 1963. Studies of nitrogen fixing bacteria. Proc. Linn. Soc. New South Wales 88:379–385. Thompson, J. P., Skerman, V. B. D. 1979. Azotobacteraceae: The taxonomy and ecology of the aerobic nitrogenfixing bacteria. Academic Press. London. Wiegel, J., Schlegel, H. G. 1984. Genus Xanthobacter. 325– 333. Bergey’s manual of systematic bacteriology. Williams and Wilkins. Baltimore. Wiegel, J., Wilke, D., Baumgarten, J., Opitz, R., Schlegel, H. 1978. Transfer of the nitrogen-fixing hydrogen bacterium Corynebacterium autotrophicum Baumgarten et al. to Xanthobacter gen. nov. Int. J. Syst. Bacteriol. 28:573–581.

Prokaryotes (2006) 5:758–777 DOI: 10.1007/0-387-30745-1_35

CHAPTER 3.2.11 ehT

areneG

x i rhtotpeL

dna

su l i toreahpS

The Genera Leptothrix and Sphaerotilus STEFAN SPRING

Introduction Representatives of the genera Leptothrix and Sphaerotilus were among the first microorganisms to be recognized in the environment and described in detail by scientists. The type species of the genus Leptothrix, L. ochracea, was already observed in the late eighteenth century and described by Roth (1797) under the synonym “Conferva ochracea.” Later, Kü tzing (1843) proposed to place this species within the genus Leptothrix. Ten years earlier, the same author had published a description of the species Sphaerotilus natans (Kü tzing, 1833), which is today still known under this name. These early publications were probably evoked by the observation of ocherous deposits (clearly visible to the naked eye) in ponds or slowly running water. A microscopic examination of these suspicious aggregates led then to the discovery of filamentous microorganisms, which were obviously responsible for the deposition of iron or ferromanganese oxides in a slimy matrix, resulting in the typical color of the formed amorphous flocs or surface films. Further investigations of the SphaerotilusLeptothrix group were induced by their potential economic importance. The massive growth of these microorganisms at industrial sites can lead to technological problems like clogging of water distribution systems or the bulking of activated sludge (Dondero, 1975). More recently, the significance of Leptothrix species in the corrosion of steel could be demonstrated (Olesen et al., 2000; Rao et al., 2000). On the other hand, a potential application of these microorganisms in the biological clearance of heavy metals from contaminated water supplies is being discussed (Nelson et al., 1999; Solisio et al., 2000). Both genera, Leptothrix and Sphaerotilus, have been always considered as closely related because of the conformity of some suspicious morphologic traits (Mulder and Van Veen, 1963). Later, their close relationship was confirmed by phylogenetic and phenotypic data, thereby justifying their treatment as a group. Characteristic traits, which distinguish members of this group

from other phylogenetically related species, are the capability to form tubular sheaths and the precipitation of copious amounts of oxidized iron or manganese.

Phylogeny and Taxonomy Phylogeny and Related Genera Members of the genera Leptothrix and Sphaerotilus comprise a phylogenetically coherent cluster within the β1-subgroup of the Proteobacteria. The similarity values among 16S rRNA gene sequences representing strains of the Sphaerotilus-Leptothrix group are in the range between 96.3 and 99.8% (Pellegrin et al., 1999). It has to be noted, however, that to date no 16S rRNA sequence of L. ochracea (the type species of the genus Leptothrix) is available. In addition, the physiology of this species is largely unknown, because it could not be isolated in pure culture. Hence, it cannot be excluded that the type species of Leptothrix is phylogenetically only distantly related to the other species of this genus. Representatives of the genera Aquabacterium, Ideonella, Rubrivivax, Roseateles and two misclassified species, [Alcaligenes] latus and [Pseudomonas] saccharophila, are phylogenetically closely related to the SphaerotilusLeptothrix group. Together they form the Rubrivivax line of descent, which is tightly associated with the Comamonadaceae (Wen et al., 1999). The relative branching order of most species within the Rubrivivax group is difficult to determine, mainly because of the restricted number of varying positions available for the estimation of phylogenetic distance values. Depending on the method and database used for tree reconstruction, it may happen that representatives from other genera get intermixed with species from the Sphaerotilus-Leptothrix group. Therefore, it is difficult to decide if the capability of sheath-formation within this group evolved from a common ancestor, i.e., if it is a monophyletic trait. Despite the close phylogenetic relationship of members of the Rubrivivax branch, a differentiation is easily possible based on phenotypic

CHAPTER 3.2.11

The Genera Leptothrix and Sphaerotilus

759

Rubrivivax gelatinosus (M60682)

Ideonella dechloratans (X72724)

Aquabacterium parvum (AF035052) Aquabacterium commune (AF035054) Aquabacterium citratiphilum (AF035050)

[Pseudomonas] sacchorophila (AB021407) Leptothrix discophora (L33975) Leptothrix mobilis (X97071) Leptothrix cholodnii (X97070)

Roseateles depolymerans (AB003623) [Alcaligenes] latus (D88007)

Sphaerotilus natans (Z18534)

5%

Fig. 1. Phylogenetic tree showing the position of members of the Sphaerotilus-Leptothrix group among representatives of the Rubrivivax-branch of the β-Proteobacteria. Sheath-forming species are labeled in red, photoautotrophic species in green, and facultatively chemolithoautotrophic species in blue. The GenBank/EMBL accession number for each sequence is shown in parentheses. With the exception of Sphaerotilus natans (DSM 565) and Leptothrix cholodnii (CCM 1827), sequences of type strains were used. This tree was reconstructed using the ARB program package (Ludwig and Strunk, 1997). It is derived from a distance matrix on a selection of 16S rRNA sequences using the neighbor-joining method of Saitou and Nei (1987). Phylogenetic distances were calculated as described by Felsenstein (1982). The sequence of Escherichia coli was used as an outgroup (not shown). The bar indicates 5% estimated sequence divergence.

characteristics. It is noteworthy that the genera in this tight assemblage represent three basically different metabolic types, namely photoautotrophs (Rubrivivax), facultative chemolithoautotrophs ([Alcaligenes] latus and [Pseudomonas] saccharophila) and respiratory chemoorganotrophs (e.g., Ideonella and Sphaerotilus). In Figure 1, a representative phylogenetic tree based on 16S rRNA gene sequence data has been selected which adequately reflects the phenotypic classification within this group. Important phenotypic characteristics useful for distinguishing members of the genera Leptothrix and Sphaerotilus from other species belonging to the Rubrivivax-lineage are listed in Table 1.

Phenotypic Characteristics Common Traits Some important phenotypic traits are shared by all members of the genera Leptothrix and Sphaerotilus: Cells are straight rods, Gram-negative and motile by flagella. Growth is filamentous under natural conditions, owing to the formation of tubular sheaths that surround single and linear chains of cells. Sheaths are produced by excretion of fibrillar polymeric substances that are crosslinked to form a mesh-like fabric closely fitting to the cells. In contrast to slimes or capsules, this matrix is not in intimate contact with the cells. In aquatic habitats rich in iron and manganese, sheaths get incrusted with ferric hydroxide (Sphaerotilus) or ferromanganese oxides (Leptothrix). Without these incrustations, they appear as very thin,

almost transparent structures and are therefore not easily recognized by phase contrast microscopy. Occasionally, cells move out of the sheath or lyse and leave behind an empty space or gap within the filament, thereby facilitating the observation of the sheath. Treatment of slide preparations with ethanol may improve the visibility of sheaths under phase contrast. Further conspicuous morphological features of members of this group include the formation of slime, holdfasts, and false branching of filaments. These traits are frequently recognized in the genus Sphaerotilus, but are less common among representatives of the genus Leptothrix. The sheaths of S. natans and several Leptothrix species are surrounded by a slime layer that may be involved in the accumulation of iron hydroxide and in the oxidation of Mn(II). Holdfasts are sticky, disc-like evaginations or blebs formed at one cell pole opposite to the flagellum by which organisms attach to walls of containers, submerged plants, stones and other surfaces. Holdfast formation can be easily observed in pure cultures of Sphaerotilus natans and Leptothrix lopholea, but is absent or variable in other cultured Leptothrix species. False branching is characteristic for Sphaerotilus natans and several Leptothrix strains. It develops if a cell attaches to an existing filament and forms a new filament, thereby forming a branch. Metabolism is strictly aerobic, respiratory and chemoorganoheterotrophic. The possibility of an autotrophic or mixotrophic growth of these bacteria with ferrous iron as electron acceptor has been discussed for many years, but up to now no

69–71

Soil

69 River water

66

NR

−

−

−

+

−

Several polar

1

Roseateles

Mud

70–72

NR

+

+

+

+

−

One polar

1

Rubrivivax

Activated sludge

68

NR

+

NR

−

−

−

Several, polar or subpolar

1

Ideonella

Drinking water

Freshwater, sediment

68–71

+

− 65–66

−

NR

−

−

+

One polar; subpolar tuft

5

Leptothrixa

+

NR

−

−

−

One polar

3

Aquabacterium

Freshwater, activated sludge

69

−

−

NR

−

−

+

Subpolar tuft

1

Sphaerotilus

a

Symbols: +, present in all species; −, absent in all species; and NR, not reported. Physiological data for Leptothrix lopholea and L. ochracea are not available. From Kersters and De Ley (1984); Palleroni (1984); Mulder and Deinema (1992); Malmqvist et al. (1994); Spring et al. (1996); Kalmbach et al. (1999); and Suyama et al. (1999).

Isolation source

G + C content (mol%) Mud

−

−

Anaerobic growth NR

+

+

Autotrophic growth with H2

NR

−

−

Photoautotrophic growth

Oxidation of Mn

−

−

Carotenoid pigments

2+

−

One polar

1

[Pseudomonas] saccharophila

−

Peritrichous

1

[Alcaligenes] latus

Formation of sheaths

Flagellation

Number of species

Characteristic

Table 1. Differential characteristics of genera and misclassified species belonging to the Rubrivivax group of the β-Proteobacteria.

760 S. Spring CHAPTER 3.2.11

CHAPTER 3.2.11

definite evidence could be presented which would support this assumption. Representatives of both genera, Leptothrix and Sphaerotilus, require vitamin B12 as an essential growth factor in mineral media. A number of Leptothrix strains have been found to require in addition thiamine and biotin as growth factors (Rouf and Stokes, 1964). In broth culture, a flocculent growth is typical. Poly-β-hydroxybutyrate is stored as reserve material. The major quinone type detected is ubiquinone Q8 (>90% of total quinones). The four major components of the cellular fatty acids are cis-9 hexadecanoic acid (cis-9 16:1), hexadecanoic acid (16:0), cis-9,11 octadecanoic acid (cis-9,11 18:1) and decanoic acid (12:0). In all strains tested, the hydroxylated fatty acid 3-hydroxydecanoic acid (3-OH 10:0) can be detected, however in some cases, only in low amounts (Kämpfer, 1998). The G +C content of DNA is similar in both genera and ranges between 68 and 71 mol%. In general, chemotaxonomic features characteristic for the Sphaerotilus-Leptothrix group are very similar in other organisms belonging to the Rubrivivax group and Comamonadaceae, so that a clear differentiation of species or even genera within this phylogenetic group is hardly possible based solely on chemotaxonomic traits. Distinguishing Traits Albeit closely related, several phenotypic characteristics allow the differentiation of the genera Leptothrix and Sphaerotilus. The ability to oxidize soluble manganese (Mn2+) compounds to solid manganic (Mn4+) oxides is restricted to Leptothrix species. This trait has been originally used for differentiation because it can be easily observed in Mn2+-containing media or habitats. Further distinguishing traits, however, were only revealed by a detailed taxonomic study of cultured strains of both genera. They include the storage of polysaccharides as reserve material only by Sphaerotilus strains, size of cells, structure of the sheath surface (Leptothrix sheaths show a rough surface when viewed by electron microscopy, in contrast to the sheaths of Sphaerotilus strains which have a smooth surface), utilization of carbon sources, and the pronounced response of S. natans to an increase of organic nutrient concentration in contrast to a poor response of most Leptothrix species. A summary of phenotypic characteristics useful for the classification of both genera is presented in Table 2.

Problems in the Taxonomy of Members of the Sphaerotilus-Leptothrix Group The Proposal of Pringsheim Members belonging to the Sphaerotilus-Leptothrix group share

The Genera Leptothrix and Sphaerotilus

761

Table 2. Main morphological and physiological characteristics of the genera Leptothrix and Sphaerotilus. Characteristic Cell dimensions Width (µm) Length (µm) Flagella Monotrichous, polar Polytrichous, subpolar Structure of sheath surfaceb Reserve material Poly-β-hydroxybutyrate Polysaccharide Major fatty acidsc

Hydroxylated fatty acid Major quinone type Oxidation of Mn2+ Growth stimulation by increase of nutrient concentration Need for vitamin B12 Carbon sources used for growthe l-Alanine l-Asparagine l-Aspartate Butyrate d-Fructose d-Glucose d-Gluconate l-Ornithine

Leptothrixa

Sphaerotilus

0.6–1.5 1.5–14

1.2–2.5 1–10

+ + Rough

− + Smooth

+ − cis-9 16:1, 16:0, cis-9,11 18:1 3-OH 10:0 Q-8 + −/+d

+ + cis-9 16:1, 16:0, cis-9,11 18:1 3-OH 10:0 Q-8 − +

+

+

− − − − D D − −

+ + + + + + + +

Symbols: +, 90% or more strains are positive; −, 90% or more strains are negative; D, 11–89% of the strains are positive. a Results are based on strains available from culture collections. b Electron microscopic observation of unstained preparations. c Number of carbon atoms:number of double bonds. d Most freshly isolated strains show no pronounced response, with the exception of Leptothrix cholodnii strains. e Utilization of carbon sources was tested in GMBN medium (Richard et al., 1985; Kämpfer, 1998). From Van Veen et al. (1978); Spring et al. (1996); and Kämpfer (1998).

some important phenotypic characteristics which are easily recognized. On the other hand, distinguishing traits are sometimes difficult to determine and require the investigation of defined pure cultures. Certain strains of this group show a considerable morphological variability, depending on the respective environmental conditions, leading to the effect that virtually identical or similar strains were described under different species names. These circumstances probably led Pringsheim (Pringsheim, 1949a; Pringsheim, 1949b) to believe that all strains of this group should be placed in the genus Sphaerotilus. He also concluded that L. ochracea would be a morphological variant of S. natans

762

S. Spring

because both species lack visible deposits of manganese oxides on their sheaths. The investigations which led to these conclusions were however inadequate because they were based on only a few partly described strains or on the observation of uncultured bacteria in natural environments. Several studies in the following years clearly revealed the mistakes in Pringsheim’s nomenclature. Nevertheless, his assumptions produced a lot of confusion in the taxonomic literature on the SphaerotilusLeptothrix group between the 1950s and 70s. In that time, for instance, a variety of manganeseoxidizing strains, which probably belong to different Leptothrix species, were described under the name Sphaerotilus discophorus (e.g., Rouf and Stokes, 1964). Instability of Phenotypic Characteristics Several difficulties regarding the taxonomy of this group are due to the loss of important phenotypic traits upon cultivation (Van Veen et al., 1978). The reversible or nonreversible loss of physiological and morphological characteristics of strains in pure culture caused divergent descriptions of species, which were originally described on the basis of observations in the natural environment (e.g., L. discophora, see below). The ability to form sheaths has been irreversibly lost by several strains of Sphaerotilus natans and by most Leptothrix strains available from public culture collections, with the exception of L. cholodnii (formerly “L. discophora”) SP-6 (= LMG 8142). Gaudy and Wolfe (1961) reported that sheath formation in Spaerotilus natans is affected by the composition of the culture medium and inhibited at high concentrations of peptone. The colony morphology of freshly isolated strains corresponds to the ability of sheath formation and can be smooth (sheathless cells) or rough and filamentous (sheathforming cells). Sometimes even various colony morphologies of the same strain can be observed on one agar plate. The manganese-oxidizing activity in Leptothrix strains is independent from the formation of sheaths and more stable in most strains, but loss of this trait has also been reported. A report on the disc- or holdfast-formation of an uncultured sheathed bacterium, which was tentatively identified as L. discophora (Carlile and Dudeney, 2001), illustrates the taxonomic problems within this group. Interestingly, the formation of basal discs or holdfasts at one end of the filament (discophora means disc-bearing) was mentioned in the original description of this species by Schwers (1912), which was based on photomicrographs of cells in natural environments. In later studies, however, disc formation was never again observed in pure cultures of this

CHAPTER 3.2.11

morphotype, so that it was concluded that the presence of this trait within the genus Leptothrix was confined to the species L. lopholea (Mulder, 1989a). If the authors of the abovementioned studies have in fact described the same species, the ability to form discs would not distinguish between L. discophora and L. lopholea, this trait being more stable in the latter. To avoid this kind of confusion, descriptions of new strains should be based on freshly isolated pure cultures that were studied, if possible, under conditions most similar to natural habitat conditions. Lack of Reference Strains A major problem in the taxonomy of the genus Leptothrix is the availability of reference strains. The type species of the genus Leptothrix, L. ochracea, is not cultured and the description is based only on morphological observations. Although the description of “L. pseudo-ochracea” was based on pure cultures, no type strain was designated. On the other hand, the type strains of L. lopholea, LVMW 124, and L. cholodnii, LVMW 99, are not available from public culture collections and have apparently been lost. The strain LMG 7171 can be used as reference strain for L. cholodnii until a neotype is designated, but for L. lopholea, no other strains are available, preventing detailed taxonomic studies on this species.

Ecology Ecology of Sheath Formation The production of a sheath takes place at the expense of energy and requires synthesis and excretion of a large amount of cellular material. Therefore, it can be assumed that sheath formation was developed by microorganisms under a selective pressure and offers several ecological and nutritional advantages in the environment. A linear arrangement of single cells within a tubular sheath enables bacteria to form filaments, without actual enlargement of cell size. It has been shown that filamentous growth is an effective strategy of bacteria to exceed the size limit of particles edible by protozoa, thereby allowing them to escape from grazing (Sommaruga and Psenner, 1995). In addition, sheaths provide cells with physical protection from infection by bacteriophages (Winston and Thompson, 1979), bacterial predators (Venosa, 1975), and bacterivorous metazoa. Obviously tough sheaths impregnated with ferric oxides cannot be consumed by metazoa and are not penetrated by bdellovibrios or phages. Rigid sheaths in combination with extracellular slime represent an ideal matrix for the rapid

CHAPTER 3.2.11

The Genera Leptothrix and Sphaerotilus

build-up of biofilms in the form of floating aggregates or dense layers on solid surfaces. The ecological advantages of biofilm formation for bacteria are well known and include reduction of flow rates in running waters, adsorption of nutrients, and establishment of heterogenous microniches characterized by various chemical gradients. The reversible inhibition of sheath formation by high nutrient concentrations in some Sphaerotilus strains may be an indication for the important role played by the sheath surface in the accumulation of nutrients (Gaudy and Wolfe, 1961). The sheath provides cells with an additional compartment close to their outer surface. Enzymes and proteins which are secreted by the cell can be incorporated in the sheath matrix, thereby allowing the extracellular production or degradation of compounds. It is possible that members of both genera use the sheath or the space between sheath and outer cell surface for the oxidation of iron and/or manganese to avoid toxic concentrations of metal compounds within the cell.

Methods for Studying Distribution and Abundance Members of the Sphaerotilus-Leptothrix group are not only abundant in natural environments, but also may play a role in industrial processes leading to several technical problems. They were associated with bulking of activated sludge (Wagner et al., 1994), corrosion of stainless steel (Olesen et al., 2000), clogging of water distribution systems (Dondero, 1975), and slime formation in paper mill factories (Pellegrin et al., 1999). To prevent damage caused by the prolific growth of these organisms, it could be advantageous to

763

monitor them in environmental samples without time-consuming enrichment and cultivation steps. Although in some samples the affiliation of bacteria to the Sphaerotilus-Leptothrix group may be assigned on the basis of phase-contrast microscopic appearance, distinction between genera or single species is hardly possible. Consequently, culture-independent methods for the identification and detection of these species were developed. Two approaches could be promising: whole cell hybridization with fluorescently labeled oligonucleotide probes and polymerase chain reaction (PCR) detection of Leptothrix species using primers targeting mofA, an enzyme gene involved in manganese oxidation. Whole Cell Hybridization Fluorescence in situ hybridization (FISH) using oligonucleotide probes is a well established technique for the identification and in situ detection of microorganisms based on signature regions of their 16S rRNA genes (Amann et al., 1995). Several probes were developed targeting 16S rRNA sequences of members of the SphaerotilusLeptothrix group. A check of published probe sequences against a current database of 16S rRNA gene sequences (Ludwig and Strunk, 1997) revealed, however, that some probes which were originally intended to be specific for a single species are in fact targeting a variety of different species, partly not even belonging to the Sphaerotilus-Leptothrix group. To minimize the risk of mistaken identification of uncultured bacteria by crossreactivity of probes, it is advisable to use simultaneously at least two different specific probes, distinguishable by fluorescence label. Suitable combinations of probes for the Sphaerotilus-Leptothrix group can be found in Table 3.

Table 3. Specificity of oligonucleotide probes targeting 16S rRNA sequences of members of the Sphaerotilus-Leptothrix group. Hybridization with probea

Sequence of: Species Leptothrix cholodnii

Strain

Accession no.

CCM 1827 X97070 SP-6 L33974 L. discophora SS-1T L33975 L. mobilis Feox-1T X97071 Sphaerotilus sp. IF4 AF072914 S. natans 6T L33980 Target region of probe (E. coli positions) Sequences with complementary target region, not affiliated to the Sphaerotilus-Leptothrix groupd

SNAb

PSP-6c

LDIb

PS-1c

+ + − + + + 656–673 13

+ + − + − − 138–155 9

− − + − − − 649–666 4

+ + + − − − 66–82 0

Symbols: +, hybridization of probe; −, no hybridization of probe under stringent conditions; and T, type strain. a Reactivity of the probe was either checked experimentally or determined by sequence comparison. b Data from Wagner et al. (1994). c Data from Siering and Ghiorse (1997). d Only sequences of cultured strains available in public data bases were counted.

764

S. Spring

CHAPTER 3.2.11

The identification of distinct species by FISH is still difficult, even by using a combination of specific probes, but the combined observation of morphological features (e.g., sheath formation) and hybridization signals should at least enable the identification of uncultured bacteria at the genus level. In Figure 2, the in situ detection of uncultured Leptothrix bacteria in an environmental wetland sample is shown. Low signal intensities of labeled cells and samples with low numbers of target cells represent frequently encountered problems of the FISH method. In habitats with low numbers of sheathed bacteria, e.g., water distribution systems or sediments, the application of microscope slides can be a promising approach to obtain suitable samples for hybridization experiments. Microscope slides were successfully used for the enrichment of sheathed bacteria by employing to advantage their ability to adhere efficiently to smooth surfaces (Spring et al., 1996; Carlile and Dudeney, 2001). After removal and cleaning, the overgrown side of these slides can be used for FISH experiments. To improve signal intensities of labeled cells, it may help to treat samples prior to hybridization with diluted hydrochloric or oxalic acid, to remove iron and manganese oxides from sheaths, which could otherwise pre-

a

vent efficient penetration of oligonucleotide probes into cells. Diagnostic PCR An alternative approach for the detection of Leptothrix bacteria was developed by Siering and Ghiorse (1997b). They used specific PCR primers for the amplification of the gene mofA, encoding a putative multi-copper oxidase involved in manganese oxidation by Leptothrix species (see section “Oxidation of Fe2+ and Mn2+” in this Chapter). The gene mofA was originally retrieved by cloning a DNA fragment of Leptothrix discophora SS-1, but homologous genes were identified also in several other Leptothrix species (Siering and Ghiorse, 1997b). The application of nondegenerate PCR primers enabled the in vitro amplification of a 706-bp portion of the gene mofA from a pure culture of L. discophora SS-1 and extracted nucleic acids from a water sample of a wetland iron seep, but not from nucleic acids extracted from a sediment sample of the same site. The obtained results were in agreement with previous observations which indicated that in this habitat, the growth of Leptothrix bacteria is restricted to ferromanganese surface films and the root zone of Lemna species in the water column. Potential advantages of this protocol are an increased specificity and sensitivity compared to the FISH method. However, in contrast to the latter method, no information on the morphology of the detected cells can be obtained.

The Genus Leptothrix Habitats

10 mm b

Fig. 2. Laser scanned epifluorescence (a) and matching laser scanned phase contrast image (b) of previously unidentified filamentous bacteria labeled by the PS-1 oligonucleotide probe in a sample of particulate material from the Sapsucker Woods wetland, Ithaca, NY. From Ghiorse et al. (1996), with permission.

Leptothrix species are widely distributed in the environment and can be easily found at sites which are characterized by a circumneutral pH, an oxygen gradient and a source of reduced iron and manganese minerals. Typical habitats include iron seeps of freshwater wetland areas, forest ponds, iron springs and the upper layers of sediments (Ghiorse and Ehrlich, 1992). At suitable sites, the massive growth of these organisms can be easily observed with the naked eye as ocherous masses emerging as surface films, solid mats or fluffy, dispersed material. Depending on the amount of oxidized manganese, the color can vary from yellowish-orange to dark brown. A wetland iron seep, shown in Fig. 3, is characterized by a pronounced ocherous color originating from a prolific growth of Leptothrix bacteria. An iron-oxidizing microbial mat that developed at an iron seep where anoxic groundwater (rich in ferrous iron) flows over a stone wall was described in detail by Emerson and Revsbech (1994). They could demonstrate that Leptothrix

CHAPTER 3.2.11

The Genera Leptothrix and Sphaerotilus

7

6

765

5 4

8 3 9

Fig. 3. Ellis Hollow iron seep (near Ithaca, NY). A wetland site with massive growth of Leptothrix species (mainly L. ochracea) resulting in typically colored surface films. From Ghiorse and Ehrlich (1992), with permission.

species represent a significant fraction of the active biomass in the upper few millimeters of this dense microbial mat. Further studies presented indirect evidence for the association of Leptothrix bacteria with lacustrine ferromanganous micronodules (Stein et al., 2001) or microbial mats in areas of possible deep-lake hydrothermal venting (Dymond et al., 1989). Most Leptothrix species are restricted to natural, unpolluted environments with low concentrations of easily degradable organic nutrients. A prominent exception is L. cholodnii, which is frequently detected in sewage sludge (Kämpfer, 1997).

Isolation Several methods for the enrichment of Leptothrix species from the environment are based on the tendency of these filamentous organisms to attach to surfaces. Mulder and Van Veen (1963) used continuous flow devices to imitate the natural growth conditions of these organisms. A schematic of such an apparatus is shown in Fig. 4. For producing iron-containing ditch water, an iron-cylinder is filled with iron-stone soil supplemented with 1 to 2 g of ferric carbonate/kg of soil. After an incubation period of about 3 weeks, during which the soil is kept saturated with water, enough ferric iron is reduced to ferrous iron to start a continuous flow of tap water percolating through the soil. The soil extract running off the iron cylinder from the upper outlet is sterilized by filtration and supplied dropwise to the upper Erlenmeyer flasks, which can be inoculated with various samples containing Leptothrix bacteria. To avoid rapid nonbiological oxidation of ferrous iron, the upper Erlenmeyer flasks can be aerated with a gas mixture of low oxygen content. Comparable to the situation in

2

1

2

10 Fig. 4. Apparatus for growing Leptothrix species in running artificial iron-containing ditch water. (1) Inlet for tap water. (2) Distributor for the incoming tap water. (3) Cylinder containing iron-stone soil. (4) Seitz filter. (5) Manifold for distributing the sterile iron-containing water. (6) Inlet for the gas mixture (1% O2, 5% CO2, and 94% N2). (7) Inoculation tube. (8) Erlenmeyer flask at a high level. (9) Erlenmeyer flask at a low level fitted with outlets (10). Reproduced with permission from Mulder and Deinema (1992).

unpolluted slowly running water, bacteria which are able to attach to solid surfaces have a selective advantage within this device because they obtain more nutrients than bacteria moving with the medium. The described apparatus can be also applied for the observation of pure cultures of Leptothrix bacteria under conditions similar to their natural environment. A less laborious enrichment method was introduced by Rouf and Stokes (1964), who filled glass cylinders with water taken from the environment and added extracted alfalfa straw as nutrient source, MnCO3, and freshly precipitated Fe(OH)3. After several days, the flocculent growth of sheathed bacteria adhering to the walls of the cylinder indicated the enrichment of Leptothrix bacteria. An alternative method, which is also based on the capability of Leptothrix species to attach to smooth surfaces, was applied by Spring et al. (1996). Microscope slides were put into a sample of freshwater sediment, and after several weeks, sheaths of filamentous bacteria encrusted with ferric oxides covered parts of the slides. The slides can be removed from the sediment and used for isolation after washing in sterile tap water to remove loosely attached sediment bacteria. Isolation of pure cultures can be achieved by streaking material from enrichment cultures on previously dried agar plates containing low levels of nitrogen and carbon sources. Enrichment cul-

766

S. Spring

CHAPTER 3.2.11

tures may not be necessary if natural environments are studied in which Leptothrix bacteria can be detected by their flocculent growth. Flocculent cell material from these sites can be washed several times with sterile tap water and streaked directly on solid media. The isolation medium used by Rouf and Stokes (1964) has the following composition (per liter of tap water): Peptone, 5.00 g; ferric ammonium citrate, 0.15 g; MgSO4 ·7 H 2O, 0.20 g; CaCl2, 0.05 g; MnSO4 ·H 2O, 0.05 g; FeCl3 ·6 H2O, 0.01 g; and agar, 12.00 g. The plates are incubated at 25°C for 5–14 days in the dark. Colonies of Leptothrix strains can be easily distinguished from most contaminating bacteria on this medium by their dark-brown color.

Identification So far six different species of Leptothrix can be distinguished on the basis of phenotypic characteristics. Reference strains for taxonomic studies are only available for three species, viz., L. cholodnii, L. discophora and L. mobilis. Descriptions of the remaining species are incomplete because they are based solely on microscopic observations (L. ochracea) or poorly characterized isolates (L. lopholea and “L. pseudoochracea”), hence preventing an accurate identification of newly isolated strains using culture-dependent methods. The name of “Leptothrix pseudo-ochracea” was not included in the Approved Lists of Bacterial Names (Skerman et al., 1980) and has therefore no formal taxonomic status. Differential characteristics of the known

Leptothrix species are listed in Table 4. Detailed morphological descriptions follow below. L. OCHRACEA Leptothrix ochracea is the most common iron-precipitating ensheathed bacterium that probably occurs all over the world in slowly running ferrous iron-containing waters, poor in readily decomposable organic matter. Low oxygen tensions seem to be required for prolific growth of this species (Emerson and Revsbech, 1994). The pronounced development and activity of L. ochracea in iron- and manganese-containing waters give rise to the accumulation and deposition of large masses of ferric oxide and, probably, manganese dioxide (MnO2), which are thought to be responsible for the formation of bog ore (see for instance, Ghiorse and Chapnick, 1983). Descriptions of this species are based on observations in the natural environment or in the laboratory on enrichment cultures of slowly running soil extract. The most typical characteristic of L. ochracea is the formation of large numbers of almost empty sheaths within a relatively short time. The mechanism of this procedure can be followed in a slide culture of the organism, in an iron-containing soil extract medium, under a phase-contrast microscope. In this way, the behavior of L. ochracea in crude culture can be observed continuously. It will be seen that chains of cells leave their sheath at the rate of 1–2 µm/ min, continuously producing a new smooth and hyaline sheath connected with the old envelope (Fig. 5a). Impregnation and covering of the sheaths with iron probably take place after the

Table 4. Differential characteristics of Leptothrix species. Characteristic Cell dimensions Width (µm) Length (µm) Flagella Monotrichous polar Polytrichous subpolar Holdfasts False branching Deposition of MnO2 on sheath surface Growth at 35°C pH 8.5 Growth ona d-Fucose Fumarate dl-Lactate G+C content (mol%)

L. ochracea

“L. pseudo-ochracea”

L. lopholea

L. cholodnii

L. discophora

L. mobilis

1.0 2–4

0.8–1.3 5–12

1.0–1.4 3–7

0.7–1.5 2.5–15

0.6–0.8 2.5–12

0.6–0.8 1.5–12

+ − − − −

+ − − − +

− + + + +

+ − − − +

+ − V + +

+ − − − ND

ND ND

ND ND

ND ND

+ +

− −

+ +

ND ND ND ND

ND ND ND ND

ND ND ND ND

− − + 68–70

+ − − 71

− + − 68

Symbols: +, present in all strains; −, absent in all strains; ND, not determined; and V, variable (depending on growth conditions). a Growth was determined in GMBN medium supplemented with the respective carbon source (Kämpfer et al., 1995).

CHAPTER 3.2.11 Fig. 5. Morphological characteristics of L. ochracea. Bar = 10 µm. (a) Phase contrast micrograph of cells moving out of sheaths and subsequently forming new sheaths. From Van Veen et al. (1978), with permission. (b) Broken old sheaths covered and impregnated with ferric hydroxide in slowly running iron(II)-containing soil extract. From Mulder and Van Veen, 1963, with permission. (c, d) Acridine orange staining of cells partly covered by a hyaline sheath. Same field viewed by differential interference contrast (c) and epifluorescence microscopy (d). Intact bacterial cells stained with acridine orange fluoresce bright green under violet illumination. Note that the chain of single cells appears as continuous filament when viewed by interference contrast. Courtesy of W. C. Ghiorse.

The Genera Leptothrix and Sphaerotilus a

c

cells have left the envelopes. Aged golden-brown and highly refractile sheaths are brittle, so that they are easily broken into relatively short fragments. Viewed under phase contrast, such fragments are very characteristic and appear similar to broken glass capillaries (Fig. 5b, c). In Mn(II)containing environments or enrichment cultures, sheaths of this species can be easily distinguished from sheaths of the other Leptothrix species which are characterized by irregular encrustations of granular MnO2. In contrast, the sheaths of L. ochracea are only impregnated with ferric oxides and lack MnO2 deposits, leading to a smooth and slender appearance of aged sheaths. The lack of MnO2 encrustations in sheaths of L. ochracea can be detected either by using an electron microscope equipped with an X-ray energy dispersive microanalysis system (Ghiorse and Ehrlich, 1992) or by treatment with oxalic acid (Carlile and Dudeney, 2001). A diluted solution (1%) of oxalic acid, a solvent for hydrated ferric oxide, can make the sheaths of L. ochracea transparent and almost disappear, whereas sheaths of other Leptothrix species are unaffected owing to the impregnation with manganese oxides. Investigators who have studied and described L. ochracea under natural conditions were unable to obtain pure cultures (Cholodny, 1926; Charlet and Schwartz, 1954). Others who thought they had isolated L. ochracea had, in fact, described one of the other species of this genus (Winogradsky, 1888; Winogradsky, 1922; Molisch, 1910; Lieske, 1919; Cataldi, 1939; Präve,

767

b

d

1957). In some instances, an organism resembling L. ochracea has been isolated, viz., “L. pseudo-ochracea.” A potential lithoautotrophic metabolism of L. ochracea with Fe(II) as electron donor could, therefore, never be proved with pure cultures and is uncertain, since the organism normally grows at a pH value of 6–7, at which Fe(II) is readily oxidized nonbiologically. In addition, its Mn (II)-oxidizing capacity, which probably occurs under natural conditions, has never been confirmed. It is, however, possible that in this species, the Mn(II) oxidizing factors are secreted into the surrounding environment leading to the precipitation of MnO2 granules away from the sheath surface. “L. PSEUDO-OCHRACEA” Cells are more slender than those of the other Leptothrix species (Table 4), and are very motile by one thin polar flagellum. Even chains of 6–10 cells may show an undulatory locomotion after leaving their sheath. This characteristic may account for the relatively large number of empty sheaths in culture, compared with the number found in cultures of most other Leptothrix species; however, L. ochracea possesses even more empty sheaths. In slowly flowing ferrous iron-containing soil extract, the sheaths become impregnated with ferric oxide and appear yellow-brown. In this respect, the organism resembles L. ochracea. However, in media with added manganese compounds, the sheaths are covered with small granules of MnO2, enabling an easy distinction from

768

S. Spring

a

the sheaths of L. ochracea which are not impregnated with manganese oxides. On Mn(II)-containing agar, the black-brown colonies are very filamentous and may exceed a width of 10 mm. On basal agar media containing 0.1% peptone and 0.1% glucose, the organism may grow in concentric rings. The normal habitat of “L. pseudo-ochracea” is the slowly running, unpolluted, iron- and manganese-containing freshwater of ditches and brooklets. This species may also be found in slightly polluted water. L. LOPHOLEA Leptothrix lopholea resembles S. natans to a greater extent than do the other Leptothrix species. It produces polytrichous subpolar flagellation and forms holdfasts and false branches. Strains may grow also in rich media. Cells usually develop short-sheathed filaments radiating from a cluster of holdfasts, giving rise to many tiny flocs when the cells are grown in liquid media (Fig. 6). Deposition of iron and manganese oxides is more pronounced on holdfasts than on filaments. On Mn(II)-containing agar media, encrustation of sheaths with MnO2 is retarded, so that colonies at first are white and later become blackbrown. Cell growth responds poorly to an increased supply of organic nutrients. Strains that do show a good response oxidize manganese more slowly. This species may be isolated from slowly flowing, unpolluted or polluted freshwater and from activated sludge. L. CHOLODNII Cells of freshly isolated strains are usually found in long chains inside the sheaths. Single motile cells may be seen outside the sheaths. In the presence of Mn(II), the

CHAPTER 3.2.11 b

Fig. 6. Leptothrix lopholea. (a) Bacterial flocs with black-brown MnO2 deposits. Reproduced with permission from Mulder and Deinema (1992). (b) Many trichomes radiating from common holdfasts. Bar = 10 µm. From Van Veen et al. (1978), with permission.

sheaths become covered with granular MnO2 (Fig. 7a). At some sheath locations, the MnO2 deposits may even exceed 10 µm. Leptothrix cholodnii, in contrast to other Leptothrix species, responds to an increased supply of organic nutrients (Table 2). This results in relatively large colonies (up to 5 mm in diameter) on nutrient-rich agar media. On Mn(II)-containing agar, blackbrown hairy colonies are formed (Fig. 7b), particularly when the organism is seeded densely. Most strains display a strong tendency to dissociate spontaneously and to produce smooth rather than the typical rough colonies (Fig. 7c). Such mutant strains are largely sheathless and oxidize manganese slightly or not at all (Mulder and Van Veen, 1963; Rouf and Stokes, 1964; Stokes and Powers, 1965). In agreement with its nutritional requirements, L. cholodnii is found in slowly running iron- and manganese-containing unpolluted waters or in polluted waters, particularly in activated sludge. L. DISCOPHORA Cells are relatively small compared to those of the other Leptothrix species described (Table 4). They may occur in narrow sheaths or be free-swimming; free cells are motile by a thin polar flagellum at one or both poles. The manganese-oxidizing and ferric oxidestoring capacities of this organism are very pronounced. In the presence of Mn(II), the sheaths are heavily but irregularly encrusted with MnO2, giving rise to sheaths of sometimes 10 µm thickness. Under natural conditions, holdfasts may be formed and sheaths are covered with a slime capsule which tapers toward the growing tip and is impregnated with hydrated ferric oxides (Carlile and Dudeney, 2001). In enrichment media with both manganese (II) and iron (II), as in slowly

CHAPTER 3.2.11 Fig. 7. Leptothrix cholodnii. (a) Sheaths encrusted with MnO2. Bar = 10 µm. Reproduced with permission from Mulder and Deinema (1992). (b, c) Colony morphologies. (b) Filamentous colony on agar with MnCO3 (medium 1, Mulder and Van Veen, 1963). Bar = 100 µm. Reproduced with permission from Mulder and Deinema (1992). (c) Smooth colony on the agar medium of Rouf and Stokes (1964). Bar = 500 µm.

The Genera Leptothrix and Sphaerotilus a

769 b

c

flowing soil extract, the sheaths become covered with a thick, dark brown, fluffy layer of ferric oxide and MnO2 which may increase the diameter of the trichomes up to about 20–25 µm (Fig. 8a). Following isolation, the ability to form sheaths is easily lost in this species (Adams and Ghiorse, 1986). False branching is regularly observed, even with sheathless strains (Fig. 8b). In older cultures, coccoid bodies and cell evaginations are formed. Colonies on the solid medium of Rouf and Stokes (1964) are about 1 mm in diameter, more or less circular in shape, flat, and darkbrown (Fig. 8c). Under certain conditions, filamentous colonies may be formed. Increasing the supply of nutrients such as glucose, peptone, methionine, purine bases, vitamin B12, biotin and thiamine only increases growth slightly. Visible aggregates are formed when grown in liquid media. The normal habitat is slowly running, unpolluted, iron- and manganese-containing water of ditches, rivers or ponds. L. MOBILIS Cells are similar in width to L. discophora cells (Fig. 9a), but are usually shorter in

length. False branching, which is typical for L. discophora, is absent in cultures of L. mobilis. Cells are highly motile by a single polar flagellum. Sheaths are not formed under laboratory conditions. Colonies on the agar medium of Rouf and Stokes (1964) are about 1 mm in diameter, circular sometimes with frayed edges, flat, smooth and dark-brown (Fig. 9b). Visible aggregates are formed when grown in liquid media. The species description is based on only one strain isolated from the sediment of a freshwater lake (Spring et al., 1996).

Preservation Stock cultures can be stored on agar slants of the medium of Rouf and Stokes (1964) for about two months at 4°C. Rouf and Stokes (1964) reported a better survival of their strains if stored at room temperature instead of refrigerated. Most Leptothrix strains do not survive lyophilization. For the long-term preservation of these strains, freezing in liquid nitrogen is recommended using suspensions of cells in freshly prepared medium supplemented with 5% dimethyl sulfoxide (DMSO) as cryoprotectant.

770

S. Spring

CHAPTER 3.2.11

a

b

Fig. 8. Leptothrix discophora. (a) Sheaths covered with ferric hydroxide and manganese dioxide in running iron(II)- and manganese(II)containing soil extract. Bar = 10 µm. From Mulder and Van Veen (1963), with permission. (b) Cells of a sheathless strain showing false branching. Bar = 10 µm. From Spring et al. (1996), with permission. (c) Smooth colony on MnSO4-containing agar. MnO2 is present within the colony and in a halo containing no bacteria. Bar = 100 µm. From Mulder and Van Veen (1963), with permission.

b

Fig. 9. Leptothrix mobilis. (a) Dimensions of single cells grown in the medium of Rouf and Stokes (1964). Bar = 10 µm. From Spring et al. (1996), with permission. (b) Colony morphology on agar medium of the same composition. Dark-brown, granular deposits of MnO2 are visible in the frayed edge of the colony. Bar = 500 µm.

c

a

Physiology The genus Leptothrix is characterized by two remarkable metabolic activities: Formation of sheaths and oxidation of iron and manganese. Oxidation of Fe2+ and Mn2+ It could be demonstrated in several studies that the deposition of metal-oxides in Leptothrix species is biologically controlled. Oxidation of manganese and iron is catalyzed by various metal-oxidizing factors actively secreted by the cell (De Vrind-de Jong et al., 1990). At circumneutral pH, biological oxidation of iron is difficult to distinguish from the chemical oxidation by oxygen. Only with the identification of an iron-oxidizing protein with a molecular

weight of 150 kDa in spent culture medium of the sheathless strain Leptothrix discophora SS-1, the capability of biological iron-oxidation in Leptothrix species could be clearly demonstrated (Corstjens et al., 1992). The function of ironoxidation is still unknown, but it seems unlikely to play a role in the generation of energy. It has to be noted, however, that Gallionella ferruginea, which is also neutrophilic and microaerophilic, thrives in the same habitats as L. ochracea and is able to gain energy from chemolithoautotrophic iron-oxidation (Hallbeck et al., 1993). Several manganese-oxidizing factors could be identified in culture supernatants of Leptothrix species. One component with an estimated molecular weight of 110 kDa could be purified and partly characterized. It is called “manga-

CHAPTER 3.2.11

nese-oxidizing factor” (MOF) and consists of protein and probably polysaccharide (Emerson and Ghiorse, 1992). It is assumed that this component is part of a larger complex that originates from membranous blebs and is associated with the sheath structure (Brouwers et al., 2000). Antibodies raised against the MOF protein of L. discophora SS-1 allowed retrieval of the gene mofA, encoding a putative multi-copper oxidase. A strong indication for an active role of this copper-dependent enzyme in manganeseoxidation is the observation that addition of small amounts of copper to actively growing cultures enhances the rates of manganese oxidation considerably (Brouwers et al., 2000). In addition to multi-copper oxidases, c-type hemes could be involved in metal-oxidation by Leptothrix species. Genes encoding proteins with potential heme-binding sites were identified as part of the operon encoding mofA. So far, the beneficial effects of manganese oxidation are largely unknown, but it is unlikely that the deposition of manganese oxides has been developed without conferring an important advantage to these bacteria. The benefits of manganese oxidation are probably so difficult to determine because they are only effective under natural conditions difficult to imitate in the laboratory. Traditionally, it was assumed that the positive effects of Mn(II)-oxidizing enzymes and manganese oxides are based on the detoxification of harmful oxygen species (superoxide and peroxide) or on the adsorption of toxic compounds (e.g., heavy metals). Recently, an interesting new perspective was introduced by Sunda and Kieber (1994). They found that manganese oxides are strong chemical oxidants able to attack complex organic compounds (e.g., humic substances), thereby leading to the release of small organic molecules which can be easily assimilated by bacteria. Structure and Composition of the Sheath The overall structure and chemical composition of sheaths formed by Leptothrix species resemble those of Sphaerotilus natans, although several distinguishing traits were found. One characteristic which can be used for the differentiation between both genera is the structure of the sheath surface. In electron micrographs of unstained preparations, the sheath surface appears rough in Leptothrix species and smooth in strains of Sphaerotilus (Figs. 10 and 12b). Emerson and Ghiorse (Emerson and Ghiorse, 1993a; Emerson and Ghiorse, 1993b) performed a detailed investigation of the ultrastructure and chemical composition of the Leptothrix sheath with the sheath-forming strain L. cholodnii (formerly “L. discophora”) SP-6. They reported that it consists of a condensed fabric of 6.5-nm-

The Genera Leptothrix and Sphaerotilus

771

Fig. 10. Electron micrograph of a Leptothrix sheath showing a rough surface. Bar = 1 µm. Reproduced with permission from Mulder and Deinema (1992).

diameter fibrils underlying a more diffuse outer capsular layer. The inner sheath layer has a thickness of 30–100 nm and seems to be associated with the outer layer of the Gram-negative cell wall by membrane evaginations. The purified sheath substance contains approximately 34% polysaccharide, 24% protein, 8% lipid, and 4% inorganic material. As major components of the polysaccharide moiety, uronic acids and amino sugars, which are probably in the N-acetylated form, were identified, whereas neutral sugars could not be detected. The sheath proteins are rich in cysteine residues (6 mol%), which confer numerous disulfide- and sulfhydryl-groups to the sheath. The heteropolysaccharide and protein moieties appear to be tightly associated or connected. On the basis of these findings, it was concluded that the principal structural elements of the sheath are proteoglycan fibrils that are covalently linked to each other by interfibril disulfide bonds resulting in a stable fabric. It is possible that the difference between the condensed inner layer and diffuse capsular layer is caused by an increased amount of free sulfhydryl groups in the exterior sheath-layer. A further characteristic of the sheath is its negative charge due to free carboxyl groups, originating mainly from the uronic acids of the sheath-proteoglycans. Both chemical groups, the free sulfhydryl and the free carboxyl groups, provide the sheath with numerous sites for binding of metal cations, especially Mn2+ and Fe2+. According to the working model

772

S. Spring

CHAPTER 3.2.11 Mn oxidizing protein(s) Exterior layer Interior layer

bleb PHA

Polyphosphate

COOS

SH

Cells

SH S S

S

Fig. 11. Schematic model of the architecture and potential function of the sheath in L. cholodnii SP-6. PHA, polyhydroxyalkanoate. From Emerson and Ghiorse (1993a), with permission.

S S

COO-

COO-

presented in Fig. 11, the anionic sheath-fibrils capture and concentrate soluble Mn(II) ions within the sheath, where they can be efficiently oxidized by excreted manganese-oxidizing factors associated with the sheath (Emerson and Ghiorse, 1993a). Manganese- and iron-oxidizing factors are probably excreted from the cell and transported to the sheath as membranous blebs containing protein complexes consisting of multi-copper oxidases and cytochromes (Emerson and Ghiorse, 1992).

The Genus Sphaerotilus Habitat Members of the genus Sphaerotilus are part of the natural microbial community in slowly running freshwater streams, ditches and ponds (Stokes, 1954). In contrast to Leptothrix sp., massive growth is typical in freshwater streams that receive high levels of organic pollution from sewage or industrial wastes, especially from paper, potato, dairy or other agricultural industries. When Sphaerotilus becomes established in a polluted river, it frequently proliferates so extensively that it lines the riverbed for long distances with a wooly, filamentous carpet (Dondero, 1975). In poor settling activated sludge (so-called “bulking sludge”), Sphaerotilus is frequently found among other filamentous microorganisms (Eikelboom, 1975). Bulking or foaming of activated sludge is a common problem in water purification plants and prevents a ready separation of bacterial sludge flocs from the treated water. It appears that conditions which stimulate the growth of Sphaerotilus natans and/or other fila-

mentous bacteria favor also the bulking of activated sludge. The correlation of conditions which favor the prolific growth of a distinct bacterial species with sludge bulking is however hardly possible owing to the high phenotypic diversity of filamentous bacteria which may be involved in this phenomenon (Kämpfer, 1997). The uncontrolled growth of Sphaerotilus in artificial environments bears several risks of economic importance. It can contribute pyrogenic material to purified water for medical injection (Dondero, 1975) and cause damage to technical equipment or machines (Pellegrin et al., 1999). On the other hand, beneficial effects of the growth of S. natans are also discussed, e.g., the biosorption of heavy metals in wastewater at low pH values (Solisio et al., 2000; Esposito et al., 2001).

Isolation Enrichment Procedures In many environments sheathed bacteria occur only in low numbers, for instance in activated sludge or nonpolluted water samples. The use of of enrichment cultures may facilitate the successful isolation of Sphaerotilus sp. from such habitats. For that purpose, variations of Winogradsky’s hay infusion technique have been used (Mulder and Deinema, 1992). Extracted alfalfa straw (Stokes, 1954) or extracted pea straw (Mulder and Van Veen, 1963) serve as the nutrient material. Most of the soluble organic matter should be removed to prevent proliferation of undesirable organisms. This can be achieved by boiling and extracting the straw after it has been cut into small pieces. Suspensions of the extracted straw in tap water (1–8% [w/v]) are distributed in Erlenm-

CHAPTER 3.2.11

eyer flasks and used as enrichment medium. After inoculation with 5–20% of a water sample or activated sludge and incubation for about one week at 22–25°C, filaments of Sphaerotilus may be seen in the medium upon microscopic observation. Often tufts of filaments can be found attached to the pieces of straw and also to the side of the flasks at or near the surface of the medium, thereby enabling easy removal using glass capillaries. Pure cultures can be obtained from homogenized portions of the enriched material as described in the following paragraph. Direct Isolation Often slimy masses of Sphaerotilus are found attached to submerged surfaces in polluted, slowly running water. Material from such sites can be successfully used for the direct isolation of S. natans or related strains by streaking on agar plates. To remove contaminating cells, the collected material should be washed with sterile tap water several times. Homogenization of the washed flocs by blending for a very short time may be useful. However, activated sludge flocs containing many filaments of Sphaerotilus should be streaked directly on the agar plates without homogenization step to avoid release of numerous, nonfilamentous cells by destroying the floc structure. The agar plates used should be dry and contain only low levels of nitrogen and carbon to limit the size of undesirable bacterial colonies, leaving large areas for the filamentous organisms. A further reason for the use of nutritionally poor media is the observation that Sphaerotilus often forms smooth colonies on nutrient-rich agar, instead of the typical rough colonies, which can be easily recognized. To inhibit the growth of contaminating fungi, the agar medium can be supplemented with cycloheximide (0.005 g ·liter –1; Pellegrin et al., 1999). A suitable isolation medium was proposed by Mulder (1989b), having the following basal composition per liter of distilled water: KH2PO4, 27 mg; K2HPO4, 40 mg; Na2HPO4 ·2 H 2O, 40 mg; CaCl2, 50 mg; MgSO4 ·7 H 2O, 75 mg; FeCl3 ·6 H2O, 10 mg; MnSO4 ·H 2O, 5 mg; ZnSO4 ·7 H 2O, 0.1 mg; CuSO4 ·5 H 2O, 0.1 mg; Na2MoO4 ·2 H2O, 0.05 mg; cyanocobalmin, 0.005 mg; peptone, 1 g; glucose, 1 g; and agar, 7.5 g. Upon inoculation and incubation of these plates at 20–30°C, colonies of Sphaerotilus may be seen and tentatively identified within a few days by their characteristically flat, dull, cottonlike appearance. Confirmation of the identification may be achieved by microscopic observation. Sphaerotilus may also be isolated by spread plate techniques with or without a previous centrifugation or homogenization step on a variety

The Genera Leptothrix and Sphaerotilus

773

of agar media (Eikelboom, 1975; Williams and Unz, 1985; Ziegler et al., 1990), including the commercially available R2A agar (Seviour et al., 1994).

Identification The genera Sphaerotilus and Leptothrix are closely related and share many phenotypic characteristics. Nevertheless, a clear differentiation of both genera is possible on the basis of phenotypic traits summarized in Table 2. The only recognized species of Sphaerotilus is currently S. natans. Typical morphological characteristics of this species are false branching of filaments, extensive slime production, attachment of filaments to solid surfaces by holdfast formation (Fig. 12a), deposition of hydrated ferric oxides on sheaths, and motility of cells by means of a bundle of subpolar flagella (Fig. 12b). In contrast to those of Leptothrix, the sheaths of Sphaerotilus occurring in natural habitats are usually thin and hyaline without encrustations by ferric or manganese oxides (Fig. 12c). However, the capability of Fe(II) oxidation can be easily demonstrated in this species by cultivation in media containing soluble iron compounds and low nutrient concentrations, e.g., soil extract enriched with ferrous iron. Under these growth conditions, the sheaths of S. natans turn yellowbrown and resemble in appearance those of Leptothrix ochracea. Most of the pronounced morphological characteristics of S. natans are largely dependent on the strain and cultivation conditions. Sheath formation can be inhibited by high levels of nutrients in the medium—peptones being more effective than carbohydrates— resulting in the formation of smooth instead of rough, filamentous colonies (Mulder and Van Veen, 1963). This loss of sheath-forming capability is reversible and can occur also spontaneously. Growth of pure cultures in liquid media is usually flocculent, but sometimes pellicular or homogenous (Pellegrin et al., 1999). Upon prolonged incubation, large, circular bodies resembling protoplasts may appear in broth cultures. Their formation is probably due to the production of enzymes involved in the decomposition of cell walls during the death phase (Phaup, 1968). The nutritional versatility of S. natans and related strains is remarkable. They can utilize a variety of carbon and nitrogen sources, tolerate a wide range of nutrient concentrations and can grow under low partial pressures of oxygen. Utilization of fructose, glucose, maltose, sucrose, lactate, pyruvate, and succinate as sole sources of carbon was reported by Stokes (1954), Mulder and Van Veen (1963), Kämpfer (1998), and Pellegrin et al. (1999). Numerous other carbon

774

S. Spring

CHAPTER 3.2.11 a

b

c

Fig. 12. Sphaerotilus natans. (a) Cells with internal granules of poly-β-hydroxybutyrate (white arrowhead) and filaments showing holdfasts (black arrowheads). Bar = 10 µm. (b) Electron micrograph of a single cell with a subpolar tuft of flagella and sheath with smooth surface. Bar = 1 µm. (c) Cells with and without a sheath and empty sheaths. Bar = 10 µm. All figures reproduced with permission from Mulder and Deinema (1992).

sources can be assimilated by strains of S. natans, but the substrate utilization patterns of strains within this species differ widely. In contrast to most Leptothrix strains, S. natans is able to assimilate relatively high concentrations of substrates from which it synthesizes considerable amounts of cellular material. Cells may contain large amounts of poly-β-hydroxybutyrate either as numerous small globules or as a few large globules (Fig. 12a). Polysaccharides may also accumulate. The synthesis of both reserve compounds is stimulated by a high carbon/nitrogen ratio in the medium or by oxygen deficiency (Mulder and Van Veen, 1963).

Preservation Stock cultures of S. natans on agar slants of the previously described media can be stored for about 3 months at 4°C. Addition of 2–3 ml of sterile tap water to the agar slants may prolong the viability for another 3 months. Preservation for longer periods is accomplished by common lyophilization techniques; however, it must be stressed that some Sphaerotilus strains do not survive lyophilization. For the long-term preservation of these strains, freezing in liquid nitrogen can be applied.

Physiology Metabolism Sphaerotilus is rarely found associated with deposits of metal oxides in natural

environments and typically thrives in habitats with normal concentrations of ferrous iron. So far, the mechanism of iron oxidation in this microorganism is poorly understood and it could be that in S. natans, iron oxidation is only a side effect of a biological reaction with a different metabolic function. Sphaerotilus natans is obligately aerobic and respiratory, but it can grow well with low concentrations of oxygen. It can readily adapt to various nutrient concentrations, growth temperatures (10–40°C) and pH values (pH 5.4–9.0), but seems to be sensitive to an increase in NaCl concentration (upper limit between 0.3 and 0.7%; Dondero, 1975). Glucose is dissimilated via the phosphogluconoate pathway and the tricarboxylic cycle. In the presence of glucose, the oxidation of other sugars, amino acids, and compounds of the tricarboxylic acid cycle seems to be repressed (Dondero, 1975). Structure and Composition of the Sheath The sheath of S. natans is, like the sheath of Leptothrix, resistant to proteases and composed of a complex of polysaccharide, protein and lipid. Romano and Peloquin (1963) detected in purified sheath material 36% carbohydrate, 28% protein and 5.2% lipid, whereas Takeda et al. (1998) obtained values of 54.1, 12.2 and 1–3%, respectively. These discrepancies may be due to the investigation of different strains of S. natans or the application of various cultivation conditions or sheath purification methods. The surface

CHAPTER 3.2.11

of the S. natans sheath is smooth and covered with an acidic exopolysaccharide composed of fucose, galactose, glucose and glucuronic acid (Gaudy and Wolfe, 1962). Slime production by exopolysaccharide secretion is much more pronounced in this species than in members of the genus Leptothrix. In contrast to the exopolysaccharide of the slime capsule, the sheath carbohydrate is free of acidic sugars and contains only glucose and galactosamine which is probably in its N-acetylated form. A heteropolysaccharide composed of glucose and galactosamine (in a molar ratio of 1 : 4) can be released from the sheath by hydrazine treatment which completely degrades the sheath structure (Takeda et al., 1998). The sheath of S. natans can be also degraded enzymatically by a kind of eliminase (produced by a Paenibacillus sp.) which attacks the polysaccharide moiety of the sheath (Takeda et al., 2000). Although the sheath structure is resistant to protease treatment, part of the sheath protein appears to be sensitive to protease attack and can be removed from the sheath by treatment with agents that reduce disulfide bonds. However, in contrast to the Leptothrix sheath, disulfide bonds apparently do not play a role in maintaining the sheath structure of S. natans. In sheaths treated with protease, the most abundant amino acids were glycine and cysteine. Only three or four other major amino acids were detected, and it was concluded that the sheath protein may consist of repeating subunits of a small peptide which is crosslinked with a polysaccharide backbone (Takeda et al., 1998). The revealed differences in the fine structure and composition of sheaths from Sphaerotilus and Leptothrix may be an indication for different functions of these structures for the respective microorganisms.

Acknowledgments. I am grateful to W. C. Ghiorse for providing unpublished photographs.

Literature Cited Adams, L. F., and W. C. Ghiorse. 1986. Physiology and ultrastructure of Leptothrix discophora SS-1. Arch. Microbiol. 145:126–135. Amann, R. I., W. Ludwig, and K.-H. Schleifer. 1995. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59:143–169. Brouwers, G. J., E. Vijgenboom, P. L. A. M. Corstjens, J. P. M. de Vrind, and E. W. de Vrind-de Jong. 2000. Bacterial Mn2+ oxidizing systems and multicopper oxidases: an overview of mechanisms and functions. Geomicrobiol. J. 17:1–24.

The Genera Leptothrix and Sphaerotilus

775

Carlile, M. J., and W. L. Dudeney. 2001. The discs of Leptothrix: Lost for 89 years. Microbiology 147:1393– 1394. Cataldi, M. S. 1939. Estudio fisió logico y sistem´tico de algunas Chlamydobacteriales (thesis). University of Buenos Aires. Buenos Aires, Argentina. 1–96. Charlet, E., and W. Schwartz. 1954. Beiträge zur Biologie der Eisenmikroben. I: Untersuchungen ü ber die Lebensweise von Leptothrix ochracea und einigen begleitenden Eisenmikroben. Schweiz. Z. Hydrol. 16:318–341. Cholodny, N. 1926. Die Eisenbakterien. Beiträge zu einer Monographie. Pflanzenforsch G. Fischer. Jena, Germany. 4:1–162. Corstjens, P. L. A. M., J. P. M. de Vrind, P. Westbroek, and E. W. de Vrind-de Jong. 1992. Enzymatic iron-oxidation by Leptothrix discophora: identification of an ironoxidizing protein. Appl. Environ. Microbiol. 58:450–454. De Vrind-de Jong, E. W., P. L. A. M. Corstjens, E. S., Kempers, P. Westbroek, and J. P. M. de Vrind. 1990. Oxidation of manganese and iron by Leptothrix discophora: use of N,N,N′,N′-tetramethyl-p-phenylenediamine as an indicator of metal oxidation. Appl. Environ. Microbiol. 56:3458–3462. Dondero, N. C. 1975. The Sphaerotilus-Leptothrix group. Ann. Rev. Microbiol. 3:77–107. Dymond, J., R. W. Collier, and M. E. Watwood. 1989. Bacterial mats from Crater Lake, Oregon and their relationship to possible deep-lake hydrothermal venting. Nature 342:673–675. Eikelboom, D. H. 1975. Filamentous organisms observed in activated sludge. Water Res. 9:365–388. Emerson, D., and W. C. Ghiorse. 1992. Isolation, cultural maintenance, and taxonomy of a sheath-forming strain of Leptothrix discophora and characterization of manganese-oxidizing activity associated with the sheath. Appl. Environ. Microbiol. 58:4001–4010. Emerson, D., and W. C. Ghiorse. 1993a. Role of disulfide bonds in maintaining the structural integrity of the sheath of Leptothrix discophora SP-6. J. Bacteriol. 175:7819–7827. Emerson, D., and W. C. Ghiorse. 1993b. Ultrastructure and chemical composition of the sheath of Leptothrix discophora SP-6. J. Bacteriol. 175:7808–7818. Emerson, D., and N. P. Revsbech. 1994. Investigation of an iron-oxidizing microbial mat community located near Aarhus, Denmark: field studies. Appl. Environ. Microbiol. 60:4022–4031. Esposito, A., F. Pagnanelli, A. Lodi, C. Solisio, and F. Veglio. 2001. Biosorption of heavy metals by Sphaerotilus natans: An equilibrium study at different pH and biomass concentrations. Hydrometallurgy 60:129–141. Felsenstein, J. 1982. Numerical methods for inferring phylogenetic trees. Q. Rev. Biol. 57:379–404. Gaudy, E., and R. S. Wolfe. 1961. Factors affecting filamentous growth of Sphaerotilus natans. Appl. Microbiol. 9:580–584. Gaudy, E., and R. S. Wolfe. 1962. Composition of an extracellular polysaccharide produced by Sphaerotilus natans. Appl. Microbiol. 10:200–205. Ghiorse, W. C., and S. D. Chapnick. 1983. Metal-depositing bacteria and the distribution of manganese and iron in swamp waters. In: R. Hallberg (Ed.) Environmental Biogeochemistry. Publ. House/FRN. Stockholm, Sweden. Ecol. Bull. 35:367–376. Ghiorse, W. C., and H. L. Ehrlich. 1992. Microbial Biomineralization of Iron and Manganese. In: R. W. Fitzpatrick

776

S. Spring

and H. C. W. Skinner (Eds.) Iron and manganese biomineralization processes in modern and ancient environments. Catena. Cremlingen-Destedt, Germany. Catena Supplement 21:75–99. Ghiorse, W. C., D. N. Miller, R. L. Sandoli, and P. L. Siering. 1996. Applications of laser scanning microscopy for analysis of aquatic microhabitats. Microsci. Res. Tech. 33:73–86. Hallbeck, L., F. Stå hl, and K. Pedersen. 1993. Phylogeny and phenotypic characterization of the stalk-forming and iron-oxidizing bacterium Gallionella ferruginea. J. Gen. Microbiol. 139:1531–1535. Kalmbach, S., W. Manz, J. Wecke, and U. Szewzyk. 1999. Aquabacterium gen. nov., with description of Aquabacterium citratiphilum sp. nov., Aquabacterium parvum sp. nov. and Aquabacterium commune sp. nov., three in situ dominant bacterial species from the Berlin drinking water system. Int. J. Syst. Bacteriol. 49:769–777. Kämpfer, P., D. Weltin, D. Hoffmeister, and W. Dott. 1995. Growth requirements of filamentous bacteria isolated from bulking and scumming sludge. Water Res. 29:1585– 1588. Kämpfer, P. 1997. Detection and cultivation of filamentous bacteria from activated sludge. FEMS Microbiol. Ecol. 23:169–181. Kämpfer, P. 1998. Some chemotaxonomic and physiological properties of the genus Sphaerotilus. Syst. Appl. Microbiol. 21:245–250. Kersters, K., and J. De Ley. 1984. Genus Alcaligenes Castellani and Chalmers 1919, 936AL. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology, 1st ed. Williams and Wilkins. Baltimore, MD. 1:361–373. Kü tzing, F. T. 1833. Beitrag zur Kenntnis ü ber die Entstehung und Metamorphose der niederen vegetalischen Organismen, nebst einer systematischen Zusammensetzung der hierher gehörigen niederen Algenformen. Linnaea 8:335–387. Kü tzing, F. T. 1843. Phycologia Generales. Leipzig, Germany. Lieske, R. 1919. Zur Ernährungsphysiologie der Eisenbakterien. Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. II 49:413–425. Ludwig, W., and O. Strunk. 1997. ARB—a software environment for sequence data. (http://www.arb-home.de/pub/ ARB/documentation/arb.ps). Malmqvist, ,ÅT. Welander, E. Moore, A. Ternström, G. Molin, G., and I.-M. Stenström. 1994. Ideonella dechloratans gen. nov., sp. nov., a new bacterium capable of growing anaerobically with chlorate as an electron acceptor. Syst. Appl. Microbiol. 17:58–64. Molisch, H. 1910. Die Eisenbakterien. G. Fischer. Jena, Germany. 1–83. Mulder, E. G., and W. L. van Veen. 1963. Investigations on the Sphaerotilus-Leptothrix group. Ant. v. Leeuwenhoek 29:121–153. Mulder, E. G. 1989a. Genus Leptothrix Kü tzing 1843, 198AL. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology, 1st ed. Williams and Wilkins. Baltimore, MD. 3:1998–2003. Mulder, E. G. 1989b. Genus Sphaerotilus Kü tzing 1833, 386AL. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology, 1st ed. Williams and Wilkins. Baltimore, MD. 3:1994– 1998.

CHAPTER 3.2.11 Mulder, E. G., and M. H. Deinema. 1992. The sheated bacteria. In: A. Balows, H.-G. Trü per, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. Berlin, Germany. 3:2612–2624. Nelson, Y. M., L. W. Lion, M. L. Shuler, and W. C. Ghiorse. 1999. Lead binding to metal oxide and organic phases of natural aquatic biofilms. Limnol. Oceanogr. 44:1715– 1729. Olesen, B. H., R. Avci, and Z. Lewandowski. 2000. Manganese dioxide as a potential cathodic reactant in corrosion of stainless steels. Corrosion Sci. 42:211–227. Palleroni, N. J. 1984. Genus I. Pseudomonas Migula 1894, 237Al. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology, 1st ed. Williams and Wilkins. Baltimore, MD. 1:141–199. Pellegrin, V., S. Juretschko, M. Wagner, and G. Cottenceau. 1999. Morphological and biochemical properties of a Sphaerotilus sp. Isolated from paper mill slimes. Appl. Environ. Microbiol. 65:156–162. Phaup, J. D. 1968. The biology of Sphaerotilus species. Water Res. 2:597–614. Präve, P. 1957. Untersuchungen ü ber die Stoffwechselphysiologie des Eisenbakteriums Leptothrix ochracea Kü tzing. Arch. Mikrobiol. 27:33–62. Pringsheim, E. G. 1949a. Iron Bacteria. Biol. Rev. Cambridge Philos. Soc. 24:200–245. Pringsheim, E. G. 1949b. The filamentous bacteria Sphaerotilus, Leptothrix, Cladothrix, and their relation to iron and manganese. Philos. Trans. R. Soc. London Ser. B. 233:453–482. Rao, T. S., T. N. Sairam, B. Viswanathan, and K. V. K. Nair. 2000. Carbon steel corrosion by iron oxidising and sulphate reducing bacteria in a freshwater cooling system. Corrosion Sci. 42:1417–1431. Richard, M. G., G. P. Shimizu, and D. Jenkins. 1985. The growth physiology of the filamentous organism Type 021N and its significance to activated sludge bulking. J. Water Pollut. Control Fed. 57:1152–1162. Romano, A. H., and J. P. Peloquin. 1963. Composition of the sheath of Sphaerotilus natans. J. Bacteriol. 86:252–258. Roth, A. W. 1797. Catalecta botanica quibus plantae novae et minus cognitae describuntur atque illistrantur. Lipsiae in Bibliopolio I.G. Mulleriano. fasc. 1. Rouf, M. A., and J. L. Stokes. 1964. Morphology, nutrition, and physiology of Sphaerotilus discophorus. Arch. Mikrobiol. 49:132–149. Saitou, N., and M. Nei. 1987. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Molec. Biol. Evol. 4:406–425. Schwers, H. 1912. Megalothrix discophora, eine neue Eisenbakterie. Zentrbl. Bakteriol. Parasitenkd. Infektkrankh. Hyg. Abt. II 33:273–276. Seviour, E. M., C. Williams, B. DeGrey, J. A. Soddell, R. J. Seviour, and K. C. Lindrea. 1994. Studies on filamentous bacteria from Australian activated sludge plants. Water Res. 28:2335–2342. Siering, P. L., and W. C. Ghiorse. 1997a. Development and application of 16S rRNA-targeted probes for detection of iron- and manganese-oxidizing sheathed bacteria in environmental samples. Appl. Environ. Microbiol. 63:644–651. Siering, P. L., and W. C. Ghiorse. 1997b. PCR detection of a putative manganese oxidation gene (mofA) in environmental samples and assessment of mofA gene homology among diverse manganese-oxidizing bacteria. Geomicrobiol. J. 14:109–125.

CHAPTER 3.2.11 Skerman, V. B. D., V. McGowan, and P. H. A. Sneath. 1980. Approved Lists of Bacterial Names. Int. J. Syst. Bacteriol. 30:225–420. Solisio, C., A. Lodi, A. Converti, and M. Del Borghi. 2000. The effect of acid pre-treatment on the biosorption of chromium(III) by Sphaerotilus natans from industrial wastewater. Water Res. 34:3171–3178. Sommaruga, R., and R. Psenner. 1995. Permanent presence of grazing-resistant bacteria in a hypertrophic lake. Appl. Environ. Microbiol. 61:3457–3459. Spring, S., P. Kämpfer, W. Ludwig, and K.-H. Schleifer. 1996. Polyphasic characterization of the genus Leptothrix: New descriptions of Leptothrix mobilis sp. nov. and Leptothrix discophora sp. nov., nom. rev. and emended description of Leptothrix cholodnii emend. Syst. Appl. Microbiol. 19:634–643. Stein, L. Y., M. T. La Duc, T. J. Grundi, and K. H. Nealson. 2001. Bacterial and archaeal populations associated with freshwater ferromanganous micronodules and sediments. Environ. Microbiol. 3:10–18. Stokes, J. L. 1954. Studies on the filamentous sheathed iron bacterium Sphaerotilus natans. J. Bacteriol. 67:278–291. Stokes, J. L., and M. T. Powers. 1965. Formation of rough and smooth strains of Sphaerotilus discophorus. Ant. v. Leeuwenhoek 31:157–164. Sunda, W. G., and D. J. Kieber. 1994. Oxidation of humic substances by manganese oxides yields low-molecularweight organic substrates. Nature 367:62–64. Suyama, T., T. Shigematsu, S. Takaichi, Y. Nodasaka, S. Fujikawa, H. Hosoya, Y. Tokiwa, T. Kanagawa, and S. Hanada. 1999. Roseateles depolymerans gen. nov., sp. nov., a new bacteriochlorophyll a-containing obligate aerobe belonging to the β-subclass of the Proteobacteria. Int. J. Syst. Bacteriol. 49:449–457. Takeda, M., F. Nakano, T. Nagase, K. Iohara, and J.-I. Koizumi. 1998. Isolation and chemical composition of the sheath of Sphaerotilus natans. Biosci. Biotechnol. Biochem. 62:1138–1143.

The Genera Leptothrix and Sphaerotilus

777

Takeda, M., K. Iohara, S. Shinmaru, I. Suzuki, and J.-I. Koizumi. 2000. Purification and properties of an enzyme capable of degrading the sheath of Sphaerotilus natans. Appl. Environ. Microbiol. 66:4998–5004. Van Veen, W. L., E. G. Mulder, and M. H. Deinema. 1978. The Sphaerotilus-Leptothrix Group of Bacteria. Microbiol. Rev. 42:329–356. Venosa, A. D. 1975. Lysis of Sphaerotilus natans swarm cells by Bdellovibrio bacteriovorus. Appl. Microbiol. 29:702– 705. Wagner, M., R. Amann, P. Kämpfer, B. Assmus, A. Hartmann, P. Hutzler, N. Springer, and K.-H. Schleifer. 1994. Identification and in situ detection of Gram-negative filamentous bacteria in activated sludge. Syst. Appl. Microbiol. 17:405–417. Wen, A., M. Fegan, C. Hayward, S. Chakraborty, and L. I. Sly. 1999. Phylogenetic relationsships among members of the Comamonadaceae, and description of Delftia acidovorans (den Dooren de Jong 1926 and Tamaoka et al 1987) gen. nov., comb. nov. Int. J. Syst. Bacteriol. 49:567– 576. Williams, M. W., and R. F. Unz. 1985. Isolation and characterization of filamentous bacteria present in bulking activated sludge. Appl. Microbiol. Biotechnol. 22:273–280. Winogradsky, S. 1888. Ü ber Eisenbakterien. Bot. Zeitschr. 46:261–270. Winogradsky, S. 1922. Eisenbakterien als Anorgoxydanten. Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. II 57:1–21. Winston, V., and T. L. Thompson. 1979. Isolation and characterization of a bacteriophage specific for Sphaerotilus natans which contains an unusual base in its deoxyribonucleic acid. Appl. Environ. Microbiol. 37: 1025–1030. Ziegler, M., M. Lange, and W. Dott. 1990. Isolation and morphological and cytological characterization of filamentous bacteria from bulking sludge. Water Res. 24:1437– 1451.

Prokaryotes (2006) 5:778–811 DOI: 10.1007/0-387-30745-1_36

CHAPTER 3.2.12 ehT

c i hpor totuaoht iL

gn i z i d i xO- a i nom A

a i re t caB

The Lithoautotrophic Ammonia-Oxidizing Bacteria HANS-PETER KOOPS, ULRIKE PURKHOLD, ANDREAS POMMERENING-RÖ SER, GABRIELE TIMMERMANN AND MICHAEL WAGNER

Introduction The lithoautotrophic ammonia-oxidizing bacteria (AOB) are well defined by their fundamental metabolism. Ammonia serves as the sole energy source and carbon dioxide is used to fulfill the carbon need. Together with the lithotrophic nitrite-oxidizing bacteria (see the chapter Nitrite Oxidizing Bacteria in this Volume), the AOB catalyze the so-called “nitrification process” (NH3 → NO2 − → NO3 − ), which has a key position in natural nitrogen cycling. Details of the biochemistry of the AOB are reported in the chapter on Oxidation of Inorganic Nitrogen Compounds as an Energy Source in Volume 2. This chapter focuses on the phylogeny, taxonomy, and diversity of AOB as well as on their distribution in nature. Winogradsky in his fundamental investigations on AOB already postulated the existence of a great diversity of species within this guild (Winogradsky, 1892; Winogradsky and Winogradsky, 1937; Winogradsky and Winogradsky, 1933). However, isolation of AOB turned out to be difficult and was neglected for several decades after Winogradsky’s classical studies. Therefore, Nitrosomonas europaea was the only species in culture for a long period. Beginning in the 1960s, S. W. Watson from the Woods Hole Research Center (Woods Hole, MA, United States) and others started a new era of isolation and culturing of novel species of AOB. Since that time, 16 species of AOB have been described and named (Watson, 1965; Watson et al., 1971c; Harms et al., 1976; Jones et al., 1988; Koops et al., 1990; Koops et al., 1991), and the existence of many other cultured species has been reported in the literature (Koops and Harms, 1985; Stehr et al., 1995a). Meanwhile, the phylogeny of the described species and the majority of the yet unnamed cultured species has become well established (Woese et al., 1984; Woese et al., 1985; Head et al., 1993; Teske et al., 1994; Pommerening-Röser et al., 1996; Purkhold et al., 2000; Purkhold et al., 2003; see the section on Phylogeny of AOB in this Chapter), providing a sufficient basis for investigations of natural AOB

populations employing classical as well as molecular techniques. In particular, the polymerase chain reaction (PCR)-assisted retrieval of 16S rRNA gene or amoA gene (encoding the active site subunit of the AOB key enzyme ammonia monooxygenase) sequences from the environment (e.g., Stephen et al., 1996; Purkhold et al., 2000) and the direct identification of AOB via fluorescence in situ hybridization FISH (e.g., Juretschko et al., 1998) allow the diversity of AOB to be monitored and their abundance in situ to be quantified. Results obtained from such investigations together with results from laboratory studies of cultured species have led to a better understanding of the ecology of this bacterial guild. The following sections describe enrichment and isolation techniques of AOB (see the section on Enrichment, Isolation, and Maintenance in this Chapter), summarize the present knowledge on their taxonomy (see the section on General Characteristics of the Genera and Species of AOB in this Chapter) and phylogeny (see the section on Phylogeny of AOB in this Chapter), and provides an overview of available techniques for investigating AOB (see the “Methods Useful for In Situ Detection of AOB” in this Chapter). The last section aims to reveal possible correlation between ecophysiological characteristics and distribution patterns of the distinct species or groups of species of AOB (see the section on Dominant Populations of AOB in Different Environments in this Chapter).

Enrichment, Isolation, and Maintenance Isolation procedures in general are targeted to the numerically most abundant AOB of the respective sample site. Therefore, the first step of enrichment should be a most probable number (MPN) dilution series in media, as far as possible reflecting the conditions of the environment under investigation. For example, in enrichments from acidic environments, low pH of the medium and addition of urea as the sole substrate for

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

energy generation is useful to isolate members of the dominant subpopulation of AOB. In general, the most important factors are the ammonia and the salt concentration in the medium. In some cases, temperature also may be of importance. Two alternative second steps on the way to isolation have been most successful. One option is additional series of dilutions, always using the highest positive dilution of the foregoing MPN series as inoculum. The other option is plating of the bacteria from the highest positive dilution of the original MPN dilution series. It is important to keep in mind that AOB sometimes grow extremely slowly. Therefore, a positive reaction in the highest dilution of an MPN series sometimes may become visible (via pH indicator) not before incubation for two or three weeks, and colony picking from agar plates should await a minimum growth period of 6–8 weeks. Standard media, useful for enrichment cultures or pure cultures, are presented in Tables 1A and 1B. For stock cultures, calcium carbonate (5 g per liter) or N-(2-hydroxyethyl)piperazine-N′2-ethanesulfonic acid (HEPES; 4.77 g per liter) can be used as a pH buffer. In working cultures, the pH should be adjusted by addition of a 10%

solution of sodium hydrogen carbonate. This addition is either performed manually (with the aid of a pH indicator) or by using an automatic pH controller. Since ammonia and not ammonium is the substrate of AOB, the pH optimum in part depends on the concentration of the added ammonium salt. Ten mM ammonium salts at a pH of about 8.0 can be regarded as good standard for sufficient growth of AOB cultures. Large volume cultures should be stirred to guarantee a sufficient supply with oxygen. In general, 30°C is a suitable growth temperature. Stock cultures should be transferred to fresh medium every 3–4 months. Storage of AOB cells in liquid nitrogen is possible, whereas freezedrying of AOB cells in general has not been successful.

General Characteristics of the Genera and Species of AOB The taxonomic framework of the AOB stems from early investigations of Winogradsky (Winogradsky, 1892; Migula, 1900; Buchanan, 1917; Winogradsky and Winogradsky, 1933). The

Table 1A. Different growth media for lithotrophic ammonia oxidizers. Medium no.b Ingredient Distilled water (ml) Seawater (ml) NH4Cl (mg) (NH4)2SO4 (mg) MgSO4 × 7H2O (mg) CaCl2 × 2H2O (mg) KH2PO4 (mg) K2HPO4 (mg) KCl (mg) NaCl (mg) Chelated iron (mg) FeSO4 × 7H2O (µg) Fe-EDTA (mg) Na2MoO4 × 2H2O (µg) (NH4)6Mo7O24 × 4H2O (µg) MnCl2 × 4H2O (µg) MnSO4 × 4H2O (µg) CoCl2 × 6H2O (µg) CuSO4 × 5H2O (µg) ZnSO4 × 7H2O (µg) H3BO3 (µg) Phenol red, 0.5% (ml) Cresol red, 0.05% (ml) a

1 1,000

2 1,000

3

130.0 200.0 20.0

4

5

1,000 1,320.0

600 400 500.0

1,000 535.0

500.0 40.0 40.0 200.0

49.3 147.0 54.4

87.0

200.0 20.0 50.0 114.0

74.4 584.0 1.0

1.0 973.1

0.5 100.0

1.0 37.1

200.0

2.0 44.6

2.0 20.0 100.0 0.1

25.0 43.1 49.4

1.0

2.0 20.0 100.0 1.0

1.0

1.0

Suitable buffers are 5.0g/l CaCO3 and 4g/l HEPES, respectively. Medium no. 1 is from Soriano and Walker (1968), for terrestrial strains. Medium no. 2 is from Watson (1971b), for terrestrial strains. Medium no. 3 is from Krü mmel and Harms (1982), for terrestrial strains. Medium no. 3 is from Watson (1965), for marine strains. Medium no. 5 is from Koops et al. (1976), for brackish-water strains.

b

779

780

H.-P. Koops et al.

CHAPTER 3.2.12

isolates originally were categorized into genera on the basis of the shape of cells. Later, the arrangement of their intracytoplasmic membranes was introduced as a second basic character. Using these criteria, the genera Nitrosomonas, Nitrosococcus, Nitrosospira, Nitrosolobus and Nitrosovibrio have been established (Watson et al., 1981; Watson et al., 1989; Koops and Möller, 1992). In the following, a brief listing of the distinguishing morphological features of the five recognized genera of AOB is given and the distinct csultured species are described. Differential characteristics of the genera of AOB and of the species of AOB are presented in Tables 2 and 3, respectively.

The Genera Genus NITROSOMONAS (Winogradsky, 1892) Cells generally are rod shaped, but sometimes spherical. Extensive intracytoplasmic membranes are arranged as peripherally located flattened vesicles. Sometimes intrusions of membranes into the protoplasm are observed (Fig. 1). Genus NITROSOSPIRA (Winogradsky and Winogradsky, 1933) Cells are tightly closed spirals and occasionally vibrio shaped. Extensive intracytoplasmic membranes are missing. Intru-

Table 1B. Different media used to ascertain that cultures of ammonia oxidizing bacteria are free of heterotrophic contaminants. Medium no.b Ingredient Distilled water Seawater Yeast extract (Difco) Peptone (Difco) Beef extract (Difco)

1

2

3

1,000ml

600ml 400ml 0.5g 0.5g 0.5g

250ml 750ml 0.5g 0.5g 0.5g

0.5g 0.5g 0.5g

a

The pH of these media should be adjusted to 7.3. Media no. 1, 2 and 3 are from Koops et al. (1976). Medium no. 1 is for terrestrial and freshwater strains, respectively; no. 2 for brackish-water strains; and no. 4 for marine strains.

b

sions of membranes into the protoplasm are sporadically observed (Fig. 2). Genus NITROSOVIBRIO (Harms et al., 1976) Cells are vibrio shaped. Extensive intracytoplasmic membranes are missing. Intrusions of membranes into the protoplasm are documented (Fig. 3). Genus NITROSOLOBUS (Watson et al., 1971c) Cells are pleomorphic lobes compartmentalized by the cytoplasmic membrane (Fig. 4). Genus NITROSOCOCCUS (Winogradsky, 1892) This genus solely represents the gamma-proteobacterial AOB. Both described species, N. oceani and N. halophilus, are characterized by large spherical to ellipsoidal cells revealing extensive intracytoplasmic membranes, arranged as a central stack of vesicles (Fig. 5).

The Cultured Species Since all AOB possess the same fundamental metabolism and morphological differences are per se limited within this group, identification on the species level often is difficult if exclusively phenotypic characters are used. The three genera Nitrosolobus, Nitrosovibrio and Nitrosospira, which phylogenetically form the Nitrosospira lineage (see the section on Phylogeny of AOB in this Chapter), are severely affected by this limitation. Nitrosomonas species within one of the distinct lineages of this genus often cannot be unambiguously identified using phenotypic characters, while species affiliated with different lineages can more easily be distinguished. In general, DNA-DNA hybridization is the method of choice to define closely related species. The above-mentioned phenotypic similarities among closely related AOB has hampered in many cases formal description of genotypically well-defined, cultured species. Below, important features of AOB species are listed.

Table 2. Characteristics of the genera of the ammonia-oxidizing bacteria. Characteristics Cell shape Cell size (µm) Flagellation of motile cells Arrangement of intracytoplasmic membranes

Nitrosococcus

Nitrosolobus

Spherical to ellipsoidal 1.5–1.8 × 1.7–2.5 Tuft of flagella

Pleomorphic lobate

Straight rods

Nitrosomonas

1.0–1.5 × 1.0–2.5 Peritrichous

Central stacks of vesicles

Compartmentalizing membranes

0.7–1.5 × 1.0–2.4 Polar to subpolar Peripheral flattened vesicles

Nitrosospira

Nitrosovibrio

Tightly coiled spirals 0.3–0.8 × 1.0–8.0 Peritrichous

Slender curved rods 0.3–0.4 × 1.1–3.0 Polar to subpolar Invaginations

Invaginations

50.6–51.4 47.9–48.5 53.8 49.3 45.6–46.0 47.9 45.6–46.0 49.4–50.0 47.4–48.0 45.7–46.3 45.5–46.1 53.5 53.9 54 50–51 50–51

G+C (mol%) − − − − − + + + + + + +/− +/− +/− + −

Carboxysomes − + + − − + − − − − − ND ND ND ND ND 42–59 ND ND ND ND ND

50–52

14–43 19–46 1.9–4.2

30–61

400 600 400 250 250 100 200 50 200 400 400 50 100 200 1000 500

Maximum ammonia tolerance NH4Cl (in mN; pH 8.0)

Symbols and Abbreviations: +, present; −, not present; +/−, present in some strains; and ND, no data.

Nitrosomonas europaea Nitrosomonas eutropha Nitrosomonas halophila Nitrosococcus mobilis Nitrosomonas communis Nitrosomonas nitrosa Nitrosomonas ureae Nitrosomonas oligotropha Nitrosomonas marina Nitrosomonas aestuarii Nitrosomonas cryotolerans Nitrosolobus multiformis Nitrosovibrio tenuis Nitrosospira briensis Nitrosococcus oceani Nitrosococcus halophilus

Species

Urease activity

Substrate (NH3) affinity (K, in µM)

Table 3. Characteristics and preferred habitats of described species of the ammonia-oxidizing bacteria.

− − + + − − − − + + + − − − + +

Salt requirement 400 400 900 500 250 300 200 150 800 600 550 200 100 250 1100 1800

Maximum salt tolerance (in mM)

Marine environments Soils (not acid) Soils, rocks and freshwater Soils, rocks and freshwater Marine environments Marine environments and salt lakes

Marine environments

Oligotrophic freshwater and natural soils

Soils (not acid) eutrophic freshwater

Sewage disposal plants, eutrophic freshwater and brackish water

Preferred habitats

CHAPTER 3.2.12 The Lithoautotrophic Ammonia-Oxidizing Bacteria 781

782

H.-P. Koops et al.

CHAPTER 3.2.12

Fig. 1A

Fig. 1B 5 mm

5 mm

0.2 mm

C

IM c

IM

0.3 mm

Fig. 1. The genus Nitrosomonas. Phase contrast photomicrographs (A) and electron micrographs of thin sections (B) of cells of different Nitrosomonas species showing the variability of shapes and sizes and the details of their ultrastructure (intracytoplasmic membranes [IM] and carboxysomes [C]).

1 mm

1 mm

3A

2A 0.5 mm

2B 3B Fig. 2. The genus Nitrosospira. Scanning electron micrograph (A) and electron micrograph of thin sections (B) of cells of Nitrosospira species.

0.3 mm

Fig. 3. The genus Nitrosovibrio. Scanning electron micrograph (A) and electron micrograph of thin sections (B) of cells of Nitrosovibrio species.

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

The Species of the Betaproteobacterial AOB The Nitrosomonas Group The cultured species of this genus can be assigned to six distinct, phylogenetically definable lineages (see the section on Phylogeny of AOB in this Chapter), comprising 11 described species. Several further genospecies are in culture but cannot sufficiently be discriminated by phenotypic properties (e.g., Koops and Harms, 1985; Stehr et al., 1995a). In

1 mm

4A

4B

0.5 mm

Fig. 4. The genus Nitrosolobus. Scanning electron micrograph (A) and electron micrograph of thin sections (B) of cells of Nitrosolobus species.

general, described species belonging to different lineages of Nitrosomonas are phenotypically well distinguishable (Table 3). However, within the distinct Nitrosomonas lineages, species discrimination in many cases is as impossible as observed for the Nitrosospira lineage. The Nitrosomonas europaea/Nc. mobilis Lineage This lineage comprises four described species, all being characterized by a relatively high salt tolerance. Two of these species, N. halophila and Nc. mobilis, have an obligatory but moderate salt requirement. All reveal relatively high affinity constants for ammonia (50– 100 µM), and all cultured strains are urease negative. Consistent with these findings the members of this group seem to prefer eutrophicated aquatic environments with heightened ionic strength. The levels of phylogenetic interrelationships among the four species of this lineage are different. While N. europaea and N. eutropha reveal absolute close relationship with each other (15–16% DNA-DNA similarity), they show only minimal DNA-DNA similarity values with N. halophila, and DNA-DNA similarities are even missing with Nc. mobilis, as measured with the S1 nuclease technique (PommereningRöser et al., 1996). Nitrosomonas europaea (Winogradsky, 1890; Watson, 1971a) Cells (0.8–1.1 × 1.0–1.7 µm) are short rods with pointed ends. Motility is not observed, and carboxysomes are missing. Often found in sewage disposal plants, cells are occasionally observed in eutrophicated freshwaters or fertilized soils. They have no obligate salt requirement, but do have a striking tolerance for

Fig. 5A Fig. 5B 0.5 mm

Fig. 5. The genus Nitrosococcus. Phase-contrast photomicrographs of whole cells (A) and electron micrographs of thin sections of cells (B) of Nitrosococcus halophilus. (C) Electron micrograph of a Nitrosococcus halophilus cell shadowed with chromium showing a tuft of flagella. (D) Freeze-etch electron micrograph of Nitrosococcus oceani showing the macromolecular arrangement of the two layers outside of the envelope.

783

5 mm

Fig. 5C Fig. 5D

1 mm

0.1 mm

784

H.-P. Koops et al.

CHAPTER 3.2.12

increasing salt concentration up to 400 mM NaCl. The G+C content of DNA is 50.6–51.4 mol%.

preferred habitat and freshwater only an occasional habitat. The G+C content of DNA is 45.6– 46.0 mol%.

Nitrosomonas eutropha (Koops et al., 1991) Cells occur singly or as short chains and are pleomorphic, rod- to pear-shaped, with one or both ends are pointed. Cells (1.0–1.3 × 1.6–2.3 µm) are motile, and carboxysomes are observed. The species is commonly distributed in sewage disposal plants, and occasionally it has been isolated from other eutrophicated environments. The species has no salt requirement but is tolerant to high salt concentration up to 400 mM NaCl. The G+C content of DNA is 47.9–48.5 mol%.

Nitrosomonas nitrosa (Koops et al., 1991) Cells (1.3–1.5 × 1.4–2.2 µm) are spheres or rods with rounded ends. Motility is not observed. Carboxysomes are present. Although eutrophicated freshwaters seem to be the preferred habitats, strains occasionally have been isolated from marine environments and a wastewater treatment plant. The G+C content of the DNA is 47.9 mol%.

Nitrosomonas halophila (Koops et al., 1991) Cells (1.1–1.5 × 1.5–2.2 µm) occasionally appear coccoid. Motile cells possess a tuft of flagella, and carboxysomes are observed. Cells have an obligate salt requirement, and some isolates are alkali tolerant up to pH 10. Cells tolerate high salt concentration (up to 900 mM NaCl). Strains were isolated from the North Sea and from soda lakes located in Mongolia (Sorokin et al., 2001). The G+C content of DNA is 53.8 mol%. Nitrosococcus (Nitrosomonas) mobilis (Koops et al., 1976) Cells (1.5–1.7 × 1.5–2.1 µm) are coccoid or rod shaped. Carboxysomes have not been observed. Motile cells possess a tuft of flagella. Cells have an obligate salt requirement. Isolates originate from the North Sea (Koops and Harms, 1985) and from an industrial wastewater treatment plant (Juretschko et al., 1998). The species seems to prefer eutrophic, aquatic environments. The G+C content of DNA is 49.3 mol%.

The Nitrosomonas oligotropha Lineage Two described species and several undescribed genospecies (e.g., Nitrosomonas sp. Nm47, Nitrosomonas sp. Nm86, Nitrosomonas sp. Nm84, and Nitrosomonas sp. Nm59) belonging to this lineage exist in culture (see the section on Phylogeny of AOB in this Chapter). The absolute majority of strains originate from oligotrophic freshwaters, and only occasional isolates are from natural, often moderately acidic (pH about 6.0) soils. All strains investigated thus far reveal strikingly low affinity constants for ammonia (few µM) and almost all isolates are urease positive. All studied strains turned out to be salt sensitive. Nitrosomonas oligotropha (Koops et al., 1991) Cells (0.8–1.2 × 1.1–2.4 µm) are rod shaped or spherical with rounded ends. Cell aggregates, obviously caused by extensive production of exopolymeric materials, are observed in the environment as well as in pure culture. Carboxysomes are missing, motility is not observed. The G+C content of DNA is 49.4–50.0 mol%.

The Nitrosomonas communis Lineage According to phenotypic properties, this lineage is divided into two sublineages. This division is also supported by DNA-DNA similarity data (Pommerening-Röser et al., 1996). One subgroup contains N. communis together with two undescribed cultured genospecies (see the section on Phylogeny of AOB in this Chapter) that are phenotypically not well distinguishable. These three species are urease negative and prefer agricultural soils with neutral pH. The other subgroup is represented by a single species, N. nitrosa. In contrast to the first subgroup, all isolates of N. nitrosa are urease positive, and aquatic environments are the preferred habitats.

The Nitrosomonas marina Lineage This lineage is represented by two described species and one undescribed genospecies (represented by the isolates Nm51 and Nm63; see the section on Phylogeny of AOB in this Chapter). All isolates originate from marine environments and reveal an obligate salt requirement. In general, isolates are urease positive.

Nitrosomonas communis (Koops et al., 1991) Cells (1.0–1.4 × 1.7–2.2 µm) are relatively large rods with rounded ends. Motility is not observed, and carboxysomes are missing. Moderately eutrophicated, pH neutral soils seem to be the

Nitrosomonas marina (Koops et al., 1991) Cells (0.7–0.9 × 1.7–2.2 µm) are slender rods with rounded ends. Motility is not observed, and carboxysomes are missing. The G+C content of DNA is 47.4–48.0 mol%.

Nitrosomonas ureae (Koops et al., 1991) Phenotypically very similar to N. oligotropha but distinguishable by a lower G+C content of the DNA (45.6–46.0 mol%).

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

Nitrosomonas aestuarii (Koops et al., 1991) Phenotypically, this species is very similar to N. marina. Cells (1.0–1.3 × 1.4–2.0 µm) are rod shaped. The G+C content of DNA is 45.7–46.3 mol%. The Nitrosomonas cryotolerans Lineage Only one species has yet been described, and only one strain is available in culture. The only cultured strain of Nitrosomonas cryotolerans (Jones et al., 1988) originates from surface water of the Kasitsna Bay (Alaska). The rod shaped cells (2.0–4.0 × 1.2–2.2 µm) requires salt and is urease positive. Carboxysomes are missing. Motility has not been observed. Intracytoplasmic membranes are not exclusively peripherally arranged, as is typical for the other members of the genus Nitrosomonas, but sometimes membranes intrude deep into the cytoplasm and often surround the nucleoplasm. Cultures can grow at temperatures as low as –5°C. The G+C content of DNA is 45.5–46.1 mol%. The Nitrosomonas sp. Nm143 Lineage In total four isolates belonging to this lineage are available in culture (Ward, 1982; Ward and Carlucci, 1985; Purkhold et al., 2003; see the section on Phylogeny of AOB in this Chapter) but were not described by name. All isolates were obtained from marine systems. The Nitrosospira Lineage Within this group of microorganisms, the three genera Nitrosospira, Nitrosolobus, Nitrosovibrio (each encompassing a single described species) and several genospecies (see the section on Phylogeny of AOB in this Chapter) have been described. While the genera show discriminative morphological characters, phenotypic species discrimination within the distinct genera is very difficult. However, the DNA G+C content differs significantly among genospecies or groups of genospecies (Koops and Harms, 1985). Different affinity constants for the binding of ammonia and different tolerances for increasing ammonia or salt concentrations could eventually serve as distinguishing features. Furthermore, temperature was observed as a factor that could be used to discriminate among species (Jiang and Bakken, 1999). However, no robust phenotypically based system for discriminating between species of the distinct genera is currently available. Unfortunately, this situation is consistent with the lack of sufficient resolution within 16S rRNA genebased phylogenetic trees of the Nitrosospira lineage (see the section on Phylogeny of AOB in this Chapter). THE GENUS NITROSOSPIRA This genus contains many cultured genospecies (see the section on

785

Phylogeny of AOB in this Chapter) that cannot sufficiently be discriminated by phenotypic characteristics (Koops and Harms, 1985). However, two groups, containing two and three distinct species, respectively, can be distinguished on the basis of DNA G+C values around 53.5 and 55.2 mol%, respectively. The existence of these two groups later has been confirmed by DNADNA hybridizations applying the S1 nuclease technique (Pommerening-Röser, 1993). Unfortunately, only a few of the numerous cultured species have yet been included in such investigations. All cultured species are more or less pleomorphic, varying in shape between tightly closed spirals and vibrio-forms. All species, thus far studied, have urease-positive as well as -negative strains at similar frequencies. Species are commonly distributed in natural soils and, occasionally, occur in freshwater environments. Via molecular studies, representatives of Nitrosospira have also been detected in marine environments (McCaig et al., 1994; Stephen et al., 1996; Phillips et al., 1999; Bano and Hollibaugh, 2000; Horz et al., 2000; Hollibaugh et al., 2002; Nicolaisen and Ramsing, 2002). However, marine isolates do not exist and salt requirement has not yet been observed among cultured Nitrosospira strains. Cells of Nitrosospira briensis (Winogradsky and Winogradsky, 1933) are motile cells and possess 1–6 peritrichous flagella. Strains of this species have been isolated from soils. The type strain is urease negative, but other isolates of this species are urease positive. The G+C content of DNA is about 54 mol%. The Genus NITROSOVIBRIO Only one species is in culture. All strains of this species have exclusively vibrio-shaped cells. Pleomorphy, as observed with several genospecies of the genus Nitrosospira, has not yet been detected. The three cultured strains of Nitrosovibrio tenuis (Harms et al., 1976) were isolated from natural soils and are urease positive. Cells are slender curved rods (0.3–0.4 × 1.1–3.0 µm). Motile cells possess 1–4 subpolar flagella. The G+C content of DNA is 53.9 mol%. The Genus NITROSOLOBUS The genus is characterized by the pleomorphic, lobate cells which are compartmentalized by the cytoplasmic membrane. One described species and one genospecies (Nitrosolobus sp. NL5) are available. Both species are distinguishable by cell size and by a significantly different DNA G+C content of 53.5 and 56.5%, respectively (Koops and Harms, 1985). Most of the isolates of both species originate from soils, generally being pH-neutral and agriculturally used. One strain of Nitrosolobus sp. was obtained from a sewage disposal plant.

786

H.-P. Koops et al.

The cells of Nitrosolobus multiformis (1.0–1.5 × 1.0–2.5 µm; Watson et al., 1971c) are pleomorphic, motile, and possess 1–20 peritrichous flagella. Cell division is by constriction. The G+C content of DNA is about 53.5 mol%.

The Species of the Gammaproteobacterial AOB The gammaproteobacterial AOB are represented by only two species of the genus Nitrosococcus: N. oceani and N. halophilus. Both species have a similar morphology. However, both are different in salt requirement, salt tolerance, and ammonia tolerance (Table 3). Moreover, the observation that all cultured strains of N. oceani possess urease whereas the two isolates of N. halophilus are urease negative indicates another discriminating character. However, more strains must be studied to verify these distinguishing features. A third species, N. nitrosus, is no longer available in culture. NITROSOCOCCUS NITROSUS (Migula, 1900) This species is no longer available in culture, and nothing is known about its ecophysiology or its phylogenetic position. This species should be placed on the list of rejected names. NITROSOCOCCUS OCEANI (Watson, 1965; Watson, 1971a; Trüper and de Clari, 1997) This species originally was described as Nitrosocystis oceanus (Watson, 1965) and later renamed “Nitrosococcus oceanus” (Watson, 1971a), before the presently accepted name Nitrosococcus oceani was established (Trü per and de Clari, 1997). Spherical or ellipsoidal cells (1.8–2.2 µm in diameter) occur singly or as pairs. Motile cells possess 1–20 flagella (tuft). Two surface layers exist, being composed of subunits arranged in rectilinear and hexagonal arrays (Fig. 5). Intracytoplasmic membranes are arranged as a stack of parallel, flattened vesicles (Murray and Watson, 1965; Watson, 1965; Watson and Remsen, 1970). All cultured strains originate from marine environments, and all are urease positive. The G+C content of DNA is between 50 and 51 mol%. NITROSOCOCCUS HALOPHILUS (Koops et al., 1990) Both species are morphologically not distinguishable from N. oceani, but differ in salt requirement and salt tolerance as well as in ammonia tolerance (Table 3). The two cultured strains were isolated from an inland salt lake and from a salt lagoon in the Mediterranean Sea, respectively. In contrast to the cultured strains of N. oceani, the two isolates of N. halophilus are urease negative. The G+C content of DNA is 50– 51 mol%.

CHAPTER 3.2.12

Phylogeny of AOB The phylogenetic framework of AOB has been established by comparative 16S rRNA gene sequence analysis of the cultured species. During the last years, this system has been significantly extended by including environmentally retrieved 16S rRNA gene sequences.

The Cultured AOB The first phylogenetic analyses of AOB were carried out by Woese and co-workers in the 1980s (Woese et al., 1984; Woese et al., 1985) and demonstrated that two phylogenetically distinct groups of AOB exist. The major group, containing the genera Nitrosomonas, Nitrosospira, Nitrosovibrio and Nitrosolobus, belongs to the class Betaproteobacteria, while the second group of AOB, represented by two species of the genus Nitrosococcus, is affiliated with the class Gammaproteobacteria (see the chapter Oxidation of Inorganic Nitrogen Compounds as an Energy Source in Volume 2). These results, obtained from 16S rRNA oligonucleotide catalogues, were later confirmed and extended via comparative 16S rRNA gene sequence analysis (Head et al., 1993; Teske et al., 1994; Utaaker et al., 1995; Purkhold et al., 2000). Today, almost complete 16S rRNA gene sequences are available for the 14 described species of AOB affiliated with the Betaproteobacteria. Furthermore, more than 100 16S-rRNA gene sequences of other AOB isolates belonging to this class have been determined. Within the Betaproteobacteria, the AOB form a well-supported monophyletic evolutionary group (Head et al., 1993; Teske et al., 1994). The minimal 16S rRNA gene sequence similarity between recognized members of this group is 89.4% (Nitrosococcus mobilis/Nitrosomonas nitrosa). Figure 6 shows a phylogenetic 16S rRNAbased tree of those AOB (15 nitrosospiras and 19 nitrosomonads) demonstrated to represent different genospecies (DNA-DNA similarity less than 60% and/or 16S rRNA sequence similarity less than 97.5%). The genera Nitrosospira, Nitrosolobus and Nitrosovibrio are closely related, and 16S rRNA phylogeny provides no convincing support for subdivision of this lineage into three genera, although morphological and ecophysiological differences exist between the different genera (see the section on General Characteristics of the Genera and Species of AOB in this Chapter). It has therefore been suggested these genera be lumped into the single genus Nitrosospira (Head et al., 1993). In contrast, the cultured nitrosomonads can be subdivided into six lineages which are consistently

CHAPTER 3.2.12 Fig. 6. 16S rRNA-based phylogenetic tree of the betaproteobacterial AOB. The tree includes only those AOB which have been demonstrated to represent different genospecies (DNADNA similarity < 60%) and for which 16S rRNA gene sequences longer than 1000 nucleotides are available. Strains with DNA-DNA similarity > 60% with each other are given in parenthesis after the respective species name. Described species are depicted in bold. Maximum likelihood, maximum parsimony, and neighbor-joining trees were calculated and merged. Multifurcations connect branches for which a relative order cannot be unambiguously determined by applying different treeing methods. Filled and empty dots indicate parsimony bootstrap values (100 resamplings) above 90% and 70%, respectively. Scale bar represents 10% estimated sequence divergence.

The Lithoautotrophic Ammonia-Oxidizing Bacteria

787

Nitrosomonas europaea Nm50 (Nm103)

Fig. 6

Nitrosomonas eutropha Nm57 Nitrosomonas halophila Nm1 (AN1-5) Nitrosococcus mobilis Nc2 (Nm104, Nm107) Nitrosomonas sp. Nm41 (Nm58) Nitrosomonas sp. Nm33 Nitrosomonas communis Nm2 Nitrosomonas nitrosa Nm90 (Nm148) Nitrosomonas sp. Nm51 (Nm63) Nitrosomonas marina Nm22 Nitrosomonas aestuarii Nm36 Nitrosomonas sp. Nm47 Nitrosomonas oligotropha Nm45 Nitrosomonas ureae Nm10 Nitrosomonas sp. Nm86 Nitrosomonas sp. Nm84 Nitrosomonas sp. Nm59 Nitrosomonas cryotolerans Nm55 Nitrosomonas sp. Nm143 Nitrosospira sp. Nsp1 (Nsp4, Nsp40, F3) Nitrosospira sp. Nsp41 Nitrosospira sp. Nv6 (Nv12) Nitrosospira briensis Nsp10 Nitrosospira sp. Nsp62 Nitrosospira sp. L115 (A4. AF) Nitrosovibrio (Nitrosospira) tenuis Nv1 Nitrosolobus (Nitrosospira) sp. NL5 Nitrosospira sp. Nsp2 Nitrosospira sp. Nsp17 Nitrosolobus (Nitrosospira) multiformis NL13 Nitrosospira sp. Nsp57 (Nsp58) Nitrosospira sp. III7 (B6) Nitrosospira sp. Nsp12 (Nsp5, III2, 40KI) Nitrosospira sp. Nsp65

retrieved using different treeing methods and which (if consisting of more than one sequence) have parsimony bootstrap support of above 90%. This phylogenetic substructure is also retrieved if all betaproteobacterial AOB isolates (56 nitrosomonads and 48 nitrosospiras), for which a 16S rRNA sequence longer than 1000 bases has been determined, are included in the treeing analysis (Figs. 7 and 8). For the different groups within the nitrosomonads, different nomenclature systems are used in the literature (Pommerening-Röser et al., 1996; Stephen, 1996; Purkhold et al., 2000; Aakra et al., 2001b; see also Fig. 7). To avoid confusion, we herein propose a hierarchical system for subdivision of the betaproteobacterial AOB as displayed below for further use. New lineages should only be postulated if 1) almost full-length 16S rRNA sequences for the respective AOB exist and 2) the lineages can be retrieved with all treeing methods with high bootstrap support. The lineages are subdivided into recognized species and genospecies with each of these units possibly

0.10

containing several strains. In the following, a brief list (consistent with the above rules) of those units within the betaproteobacterial AOB is presented: Betaproteobacterial AOB Nitrosospira lineage (Genus Nitrosospira) (Genus Nitrosolobus) (Genus Nitrosovibrio) Nitrosomonas group N. europaea/Nc. mobilis lineage N. communis lineage N. marina lineage N. oligotropha lineage N. cryotolerans lineage Nitrosomonas sp. Nm143 lineage Although 16S rRNA phylogeny of the nitrosospiras does not reveal an obvious substructure, Stephen et al. (1996) suggested subdivision of this group into three “clusters” (clusters 2, 3 and 4, Fig. 8; cluster 1 contains no cultured species,

788

H.-P. Koops et al.

Nitrosomonas sp. TK794 Nitrosomonas sp. IWT202 Nitrosomonas sp. F6 Nitrosomonas eutropha Nm57 Nitrosomonas sp. CNS326 Nitrosomonas sp. GH22 Nitrosomonas sp. G1 Nitrosomonas sp. K1 Nitrosomonas sp. F5

Fig. 7

N. europaeal Nc. mobilis lineage (Cluster 7) 92.4

CHAPTER 3.2.12

Nitrosomonas sp. HPC101 Nitrosomonas sp. DYS323 Nitrosomonas sp. DYS317 Nitrosomonas sp. WH-2 Nitrosomonas europaea Nm50 Nitrosomonas europaea Nm103 Nitrosomonas sp. ENI-11 Nitrosomonas sp. Koll-21 Nitrosomonas sp. IWT514 Nitrosomonas sp. AN1-5 Nitrosomonas halophila Nm1 Nitrosomonas sp. Nm107 Nitrosomonas sp. Nm104 Nitrosomonas mobilis Nc2 Nitrosomonas mobilis Nm93 Nitrosomonas sp. TNO632 Nitrosomonas communis Nm2 Nitrosomonas sp. Nm33 Nitrosomonas sp. Nm41 Nitrosomonas sp. Nm58 Nitrosomonas nitrosa Nm90 Nitrosomonas sp. Nm148

N. communis lineage (Cluster 8) 94.3

Nitrosomonas sp. B19-i-NH4 Nitrosomonas sp. TA 921-i-NH4

N. marina lineage (Cluster 6b) 95.6

Nitrosomonas sp. Nm63 Nitrosomonas sp. C-113 Nitrosomonas sp. Nm51 Nitrosomonas marina Nm22 Nitrosomonas sp. 122 Nitrosomonas sp. C-45 Nitrosomonas sp. NH4W Nitrosomonas sp. N03W Nitrosomonas sp. URW Nitrosomonas aestuarii Nm36 Nitrosomonas ureae Nm10 Nitrosomonas sp. AL212 Nitrosomonas sp. Nm47 Nitrosomonas oligotropha Nm45 Nitrosomonas sp. Nm86 Nitrosomonas sp. Nm59 Nitrosomonas sp. JL21 Nitrosomonas sp. Nm84 Nitrosomonas cryotolerans Nm55 Nitrosomonas sp. TT140-89A Nitrosomonas sp. TT140-098-2 Nitrosomonas sp. Nm143 Nitrosomonas sp. C-17

N. oligotropha lineage (Cluster 6a) 95.8

Nitrosomonas cryotolerans lineage Nitrosomonas sp. Nm143 lineage (Cluster 9) 97.5 Nitrosospira lineage 96.1

0.10

Fig. 7. 16S rRNA based phylogenetic tree of the nitrosomonads. The tree includes all isolates for which 16S rRNA gene sequences longer than 1000 nucleotides are available. Described species are depicted in bold. Maximum likelihood, maximum parsimony, and neighbor-joining trees were calculated and merged. Multifurcations connect branches for which a relative order cannot be unambiguously determined by applying different treeing methods. Filled and empty dots indicate parsimony bootstrap values (100 resamplings) above 90% and 70%, respectively. For each cluster, the minimum 16S rRNA sequence similarity between two of its members is depicted. Sequences included in the analysis were published by Head et al. (1993), Suwa et al. (1997), Juretschko et al. (1998), Sorokin et al. (1998), Aakra et al. (1999a), Aakra et al. (1999b), Yamagata et al. (1999), Purkhold et al. (2000), and Purkhold et al. (2002). Sequences of strains CNS326, G1, K1, IWT202, TK794, WH-2, Koll-21, DYS317, DYS323, IWT514, TNO632, marine bacteria C-45, NH4W, 122, URW, NO3W, C-113, TT140-098-2, TT14089A, and estuarine bacteria TA 921-i-NH4, B19-i-NH4, C-17 are unpublished but available at GenBank. Scale bar represents 10% estimated sequence divergence.

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria Nitrosospira sp. Nsp1 Nitrosospira sp. Nsp4 Nitrosospira sp. GS833

Fig. 8

Nitrosospira sp. E12 Nitrosospira sp. TYM9 Nitrosospira sp. F3 Nitrosospira sp. NRS527 Nitrosospira sp. Nsp40 Nitrosospira sp. Nsp41 Nitrosospira sp. Np39-19 Nitrosospira sp. Np22-21 Nitrosospira sp. Nv6 Nitrosospira sp. Nv12 Nitrosospira birensis Nsp10 Nitrosospira sp. Nsp62 Nitrosospira sp. L115

“Cluster 3”

Nitrosospira sp. 24C Nitrosospira sp. AF Nitrosospira sp. A16 Nitrosospira sp. A4 Nitrosovibrio (Nitrosospira) tenuis Nv1 Nitrosolobus (Nitrosospira) sp. NL5 Nitrosospira sp. NpAV Nitrosospira sp. C-141 Nitrosospira sp. Nsp17 Nitrosospira sp. Nsp2 Nitrosospira sp. RY3C Nitrosospira sp. RY6A Nitrosospira sp. TCH716 Nitrosospira sp. PJA1 Nitrosolobus (Nitrosospira) multiformis NL13 Nitrosospira sp. Nsp58 Nitrosospira sp. Nsp57 Nitrosospira sp. O13 Nitrosospira sp. AHB1 Nitrosospira III7 Nitrosospira sp. T7

“Cluster 2”

Nitrosospira sp. B6 Nitrosospira sp. O4 Nitrosospira sp. Ka3 Nitrosospira sp. Ka4 Nitrosomonas

“Cluster 4”

Nitrosospira sp. III2 Nitrosospira sp. D11 Nitrosospira sp. GM4

“Cluster 0”

Nitrosospira sp. 40KI Nitrosospira sp. Nsp12 Nitrosospira sp. Nsp5 Nitrosospira sp. Nsp65

0.10

Fig. 8. 16S rRNA-based phylogenetic tree of the highly related genera Nitrosospira, Nitrosolobus, and Nitrosovibrio. The tree includes all isolates for which 16S rRNA gene sequences longer than 1000 nucleotides are available. Described species are depicted in bold. Maximum likelihood, maximum parsimony, and neighbor-joining trees were calculated and merged. Multifurcations connect branches for which a relative order cannot be unambiguously determined by applying different treeing methods. Filled and empty dots indicate parsimony bootstrap values (100 resamplings) above 90% and 70%, respectively. Sequences included in the analysis were published by Head et al. (1993), Teske et al. (1994), Utaaker et al. (1995), Tokuyama et al. (1997), Aakra et al. (1999a), Aakra et al. (1999b), Aakra et al. (2001b), and Purkhold et al. (2002). Sequences of strains GS833, E12, NRS527, NpAV, RY6A, RY3C, TCH716, and PJA1 are unpublished but available at GenBank. Scale bar represents 10% estimated sequence divergence.

789

see the section on Environmental Sequences in this Chapter). Purkhold et al. (2000) subsequently extended this system by the proposing cluster 0. These nitrosospira clusters are found with all treeing methods but not all of them are well supported by bootstrap analysis. Therefore, clusters of the Nitrosospira lineage are not comparable with the distinct lineages within the Nitrosomonas group. This reflects the fact that the minimum 16S rRNA sequence similarity of 96.1% within the entire Nitrosospira group is higher than those within the individual, described Nitrosomonas lineages (Fig. 7). It should also be noted that DNA-DNA hybridization analyses suggest that the Nitrosospira clusters 0, 2 and 4 might each represent only one single species (H.-P. Koops, unpublished observation). Taken together, it is thus questionable whether subdivision of the Nitrosospira lineage into the above-mentioned clusters or other subdivisions (Aakra et al., 2001b) is justified. Owing to the limited discriminatory power of comparative 16S rRNA analysis, in particular for the Nitrosospira lineage, other phylogenetic marker molecules were tested for phylogeny inference of AOB. Aakra and co-workers determined the 16S–23S rRNA gene intergenic spacer region (ISR) of several AOB and used these sequences for phylogenetic analysis (Aakra et al., 2001b). In that study, the authors postulated highly consistent ISR- and 16S rRNA-based tree topologies for AOB and thus suggested the usefulness of the ISR for future studies on AOB diversity and evolution. However, careful inspection of the phylogenetic trees presented in that publication reveal significant topological incongruencies between 16S rRNA and ISR trees. Owing to the high variability of the ISR, these sequences are very difficult to align and not suitable to reliably determine evolutionary relationships above the species level. Recently, the amoA gene coding for the active site polypeptide of the ammonia monooxygenase has been used as an additional phylogenetic marker molecule for AOB (McTavish et al., 1993; Klotz and Norton, 1995; Rotthauwe et al., 1995; Suwa et al., 1997; Hommes et al., 1998; Alzerreca et al., 1999; Yamagata et al., 1999; Horz et al., 2000; Purkhold et al., 2000; Aakra et al., 2001a; Casciotti and Ward, 2001; Purkhold et al., 2003). PCR primers that allow amplification of a 453bp fragment of this gene are generally used in these studies (Rotthauwe et al., 1997; modified by Stephen et al., 1999). Phylogeny inference based on the deduced amino acid sequence of the amoA gene fragment is overall consistent with the 16S rRNA phylogeny of AOB. Members of the genera Nitrosospira, Nitrosolobus and Nitrosovibrio form a tight monophyletic grouping with no obvious substructure. Within the

790

Fig. 9

H.-P. Koops et al.

CHAPTER 3.2.12

Nitrosomonas eutropha Nm57 Nitrosomonas sp. TK794 Nitrosomonas sp. F6, F5, GH22 Nitrosomonas sp. ENI-11 Nitrosomonas europaea Nm50

N. europaea/ Nc. mobilis lineage (Cluster 7)

Nitrosomonas halophila Nm1 Nitrosomonas sp. AN1 Nitrosomonas mobilis Nc2 Nitrosomonas sp. Nm41 Nitrosomonas sp. Nm58

N. communis lineage (Cluster 8)

Nitrosomonas communis Nm2 Nitrosomonas sp. Nm33 Nitrosomonas nitrosa Nm90 Nitrosomonas sp. Nm148

Nitrosomonas sp. Nm51 Nitrosomonas marina Nm22

N. marina lineage (Cluster 6b)

Nitrosomonas aestuarii Nm36 Nitrosomonas ureae Nm10 Nitrosomonas sp. AL212 Nitrosomonas sp. JL21 Nitrosomonas sp. Nm59 Nitrosomonas sp. Nm84

N. oligotropha lineage (Cluster 6a)

Nitrosomonas sp. Nm86 Nitrosomonas oligotropha Nm45 Nitrosomonas sp. Nm47 Nitrosomonas sp. Nm143

Nitrosomonas sp. Nm143 lineage

Nitrosomonas cryotolerans Nm55

N. cryotolerans lineage

Nitrosospira lineage

nitrosomonads the N. europaea/Nc. mobilis lineage and the N. marina lineage are also found with all treeing methods, while the N. communis and the N. oligotropha lineage are not always monophyletic (Fig. 9). If amoA nucleic acid sequences are used for phylogenetic analysis, basically the same picture emerges with the exception that the N. europaea/eutropha lineage is no longer monophyletic (data not shown). If compared to the 16S rRNA-based phylogeny of AOB, AmoA analysis does provide less resolution, reflecting that a relatively short (151 positions) and highly conserved (93 positions have an identical amino acid in at least 98% of the betaproteobacterial AOB) amino acid sequence stretch is used as marker. The information content of AmoA sequences could be significantly extended in future studies by the development of primers that allow the amplification of more complete amoA gene fragments. First attempts

Fig. 9. AmoA-based phylogenetic tree of the betaproteobacterial AOB. Described species are depicted in bold. The 453-bp gene fragment obtainable with the most commonly used amoA PCR primers (Rotthauwe et al., 1995) was used for phylogeny inference. AmoA sequences shorter than 414 nucleotides were excluded from the analysis. Protein maximum likelihood, protein maximum parsimony, neighbor-joining, and Fitch trees were calculated and merged. Multifurcations connect branches for which a relative order cannot be unambiguously determined by applying different treeing methods. Filled and empty dots indicate parsimony bootstrap values (100 resamplings) above 90% and 70%, respectively. Sequences included in the analysis were published by McTavish et al. (1993), Holmes et al. (1995), Rotthauwe et al. (1995), Suwa et al. (1997), Yamagata et al. (1999), Aakra et al. (2001a), Casciotti et al. (2001), Sorokin et al. (2001), Norton et al. (2002), and Purkhold et al. (2003). Sequences of Nitrosospira sp. C-57 and Nitrosomonas sp. TK794 are unpublished but available at GenBank. Scale bar represents 10% estimated sequence divergence.

0.10

in this direction were recently published by Norton et al. (2002). Correlation plots of amoA and AmoA similarity versus 16S rRNA similarity of all possible pairs of betaproteobacterial AOB isolates (Fig. 10) demonstrate that 1) 16S rRNA is more conserved than amoA or AmoA and 2) AOB showing below 80% amoA nucleic acid similarity (or 85% AmoA amino acid similarity) always possess less than 97.0% 16S rRNA sequence similarity (the currently accepted 16S rRNA threshold value for bacteria genospecies assignment; Stackebrandt and Goebel, 1994). AmoA sequences of a new AOB isolate with less than 80% nucleic acid similarity (or 85% amino acid similarity) to described AOB species are thus indicative of a previously undiscovered species. An amoA or AmoA sequence with a higher similarity to a described AOB species can represent multiple gene copies, different strains

Fig. 10. Correlation plots of 16S rRNA similarity with amoA (panel A) and AmoA (panel B) similarity values. Solid lines indicate 16S rRNA threshold values for species delineation. Dotted lines indicate the amoA and AmoA threshold values below which amoA and AmoA sequences are indicative of novel AOB species.

The Lithoautotrophic Ammonia-Oxidizing Bacteria

% 16S rRNA gene similarity

CHAPTER 3.2.12

100 99 98 97 96 95 94 93 92 91 90 89 88 87 86

791

multiple copies of the same strain different strains of the same species different species

55

60

65

70

75

80

85

90

95 100

% 16S rRNA gene similarity

% amoA similarity

100 99 98 97 96 95 94 93 92 91 90 89 88 87 86

multiple copies of the same strain different strains of the same species different species

55

60

65

70

75

80

85

90

95 100

% AmoA similarity

of this species, or a novel AOB species. The latter possibility exists since 16S rRNA similarities between different species can be higher than 97%.

Environmental Sequences Molecular diversity surveys of AOB almost exclusively focussed on betaproteobacterial AOB. Stephen et al. (1996) were the first to investigate the diversity of these bacteria in an environmental sample using directly retrieved 16S rRNA gene sequences (see the section on Molecular Techniques in this Chapter). Since then, AOB species richness of numerous natural and engineered systems has been analyzed by the 16S rRNA approach. However, the PCR primers applied in several of these studies do not cover all recognized AOB and thus probably provided a biased view of the natural AOB diversity and their environmental distribution (see the

section on Molecular Techniques in this Chapter). Interestingly, of the 508 16S-rRNA clones affiliated with betaproteobacterial AOB (status August 2002), more than 83% cluster with cultured representatives of this guild. The remaining clones form two novel clusters, one within the nitrosomonads (environmental lineage 5) and one within the nitrosospiras (cluster 1; Fig. 11). On the basis of the results of the various AOB 16S rRNA diversity surveys, environmental distribution patterns of the different AOB clusters can be revealed and compared to the findings obtained with cultivation-based methods (Fig. 12). A more detailed discussion of environmental distribution patterns of the different AOB is provided in a separate section (see the section on Distribution of AOB in Nature in this Chapter). The natural diversity of ammonia oxidizers in various environments also has extensively been studied by comparative sequence analysis

792

H.-P. Koops et al.

CHAPTER 3.2.12

Isolates Enrichments 24 59

Nitrosospira

43

“Cluster 0”

6

-

18

“Cluster 1”

-

-

59

“Cluster 2”

6

2

35

“Cluster 3”

31

23

118

“Cluster 4”

3

1

30

2

-

-

1 -

8

16

8

12

92

2 4 6

6 1 2

4 1 2

5

1

65

14

3

50

172

34 2 14 112

Nitrosospira sp. Nsp57 + Nsp58 Nitrosospira sp. Nsp65 environmental clones N. oligotropha, N. ureae + RS

10

Nitrosospira sp. + RS N. marina + RS N. aestuarii + RS

71

N. europaea + RS

12 6

Nitrosomonas

Environ clones

67

N. eutropha + RS

env. lineage 5 N. oligotropha lineage

N. marina lineage

N. europaeal Nc. mobilis lineage

6

N. halophila + RS

6

-

2

12

N. mobilis + RS

5

2

5

7

N. communis + RS

6

-

2

21

Nitrosomonas sp. Nm143 + RS

4

8

9

1

-

-

110

69

508

Nitrosomonas cryotolerans

Â

Fig. 11. Schematic 16S rRNA-based phylogenetic classification of the betaproteobacterial AOB. Multifurcations connect branches for which a relative order could not be unambiguously determined by applying different treeing methods. The height of each triangle represents the number of sequences in the cluster. It should be noted that 16S rRNA sequences shorter than 1000 nucleotides were only considered if they could be unambiguously assigned to one of the lineages. All other short 16S rRNA sequences had to be omitted because their phylogenetic analysis cannot reliably be performed (Ludwig et al., 1998). The 16S rRNA sequences used in this analysis were published by Takahashi et al. (1992), Head et al. (1993), McCaig et al. (1994), McCaig et al. (1999), Teske et al. (1994), Rotthauwe et al. (1995), Utaaker et al. (1995), Pedersen et al. (1996), Stephen et al. (1996), Kowalchuk et al. (1997), Kowalchuk et al. (1998), Kowalchuk et al. (2000a), Kowalchuk et al. (2000b), Suwa et al. (1997), Tokuyama et al. (1997), Juretschko et al. (1998), Logemann et al. (1998), Princic et al. (1998), Speksnijder et al. (1998), Aakra et al. (1999a), Aakra et al. (1999b), Aakra et al. (2000), Aakra et al. (2001b), Bruns et al. (1999), Mendum et al. (1999), Philips et al. (1999), Philips et al. (2000), Radeva et al. (1999), Whitby et al. (1999), Whitby et al. (2001a), Whitby et al. (2001b), Yamagata et al. (1999), Abd El Haleem et al. (2000), Bano et al. (2000), Ward et al. (2000), Bollmann et al. (2001), Burrell et al. (2001), Chang et al. (2001), de Bie et al. (2001), Daims et al. (2001a), Gieseke et al. (2001), Smith et al. (2001), Sorokin et al. (2001), Hollibaugh et al. (2002), Nicolaisen et al. (2002), Regan et al. (2002), and Purkhold et al. (2003). In addition, unpublished 16S rRNA sequences (Accession numbers: AF107527, AJ441258, AJ441259, AJ441260, AJ441261. AJ441262, AJ441263, AJ441264, AJ441265, AJ441266] AJ441267, AJ441268, AJ441269, AJ441270, AJ441271, AJ441272], AJ441273, AJ441274, AJ441275, AJ441276, AJ441277, AJ441278, AJ441279, AJ441280, AJ441281, AJ441282, AJ441283, AJ441284, AJ441285, AF034139, AF034140, AF034141], AF034142, AF034143, AF034144, AF034147, AJ318197, U57617, AF510862, AF510863, AF510864, AF510865, AJ245751, AJ245752, AJ245753, AJ245754, AJ245755, AJ245756, AJ245757, AJ245758, AJ245759, AJ245760, AJ431350, AJ431351], AF414581, Y10128, Y10127, AJ224941, AF353155, AF353156, AF353157, AF353158, AF353159, AF353161, AF353162, AF353163, AF353164, AF359341, AF363287, AF363289, AF363290, AF363291, AF363292, AF363293, and A036898) available in GenBank were analyzed. RS: related sequences.

of environmentally retrieved amoA clones (Rotthauwe et al., 1997; Helmer et al., 1999; Stephen et al., 1999; Baribeau et al., 2000; Horz et al., 2000; Ivanova et al., 2000; Nold et al., 2000; Purkhold et al., 2000; Schmid et al., 2000; Chang et al., 2001; Gieseke et al., 2001; Oved et

al., 2001; Dionisi et al., 2002; Sakano et al., 2002). In total, 383 amoA clones are currently available from these studies (status August 2002). Consistent with the 16S rRNA-based AOB diversity surveys, most amoA clones obtained are affiliated with sequence clusters

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

Nitrosospira “Cluster 0”

Clones/Enr. (4 habitats)

Nitrosospira “Cluster 1”

Clones/Enr. (8 habitats)

Nitrosospira “Cluster 2”

Clones/Enr. (10 habitats)

Nitrosospira “Cluster 3”

Clones/Enr. (26 habitats)

Nitrosospira “Cluster 4”

Clones/Enr. (10 habitats)

enironmental lineage 5

Clones/Enr. (7 habitats)

N. oligtropha lineage

Clones/Enr. (15 habitats)

N. marina lineage

Clones/Enr. (9 habitats)

N. europaeal Nc. mobilis lineage

Clones/Enr. (18 habitats)

N. communis lineage Nitrosomonas sp. Nm143 lineage

793

Isolates (6 habitats)

Isolates (0)

Isolates (3 habitats)

Isolates (20 habitats)

Isolates (3 habitats)

Isolates (0)

Isolates (9 habitats)

Isolates (10 habitats)

Isolates (12 habitats)

Isolates (6 habitats) Clones/Enr. (1 habitats) Isolates (4 habitats) Clones/Enr. (6 habitats)

0%

10%

20%

30%

40%

50%

60%

70%

80%

soil (n=34, m=19)

marine environment (n=13, m=8)

wastewater treatment plant (n=15, m=6)

coastal sand dune (n=0, m=2)

freshwater environment (n=4, m=10)

estuary (n=3, m=2)

90%

100%

others (n=4, m=3)

Fig. 12. Environmental distribution patterns of AOB clusters. For each cluster, the origin (sampling site) of each 16S rRNA sequence belonging to this cluster was determined. Subsequently, we counted the number of different sampling sites in which members of a particular cluster were detected. This number of habitats, which is given in parentheses, was set to 100%. In the next step, the sampling sites were grouped into the categories wastewater treatment plants, soil (including a rhizoremediation plant), freshwater (including sediment), marine (including sediment), estuary, and coastal sand dune. All other habitats were lumped together. For each cluster the proportion of each category is shown. For comparison, an identical analysis was performed of the available isolates for each cluster. The displayed legend explains the color-coding of the habitats. In addition, the number of different samples of a particular habitat category analyzed by the 16S rRNA approach (n) or cultivation (m) is given. For example, 16S rRNA AOB clones are available from 34 different soil samples and 19 soils were examined by AOB cultivation.

defined by cultured ammonia oxidizers (Table 4). The environmental distribution patterns of AOB inferred from AmoA analysis (Fig. 13) confirm the results from the 16S rRNA studies (Fig. 12), although the limited resolution provided by AmoA does not allow specific data for the individual Nitrosospira clusters to be extracted or the N. oligotropha to be distinguished from the N. marina cluster. According to the threshold values inferred from the amoA/ 16S rRNA correlation plots (Fig. 10), only two (clone GLII-9 from a wastewater treatment plant [wwtp] and clone Pluß see from a lake) of the environmental amoA clones probably represent novel AOB species. These observations confirm that ammonia oxidizers already deposited in culture collections might be surprisingly representative of the natural diversity within this guild. However, it is important to stress that the sequences from 16S rRNA or amoA gene libraries from environmental samples rarely

contain sequences completely identical to those of cultured organisms. Since no DNA-DNA hybridization data can be obtained for noncultured organisms, it can, according to the current species definition in bacteriology (Stackebrandt et al., 1988), not be decided whether these sequences represent novel or known ammonia oxidizer species. In this context, one should however bear in mind that PCR and cloning procedures introduce microvariation artifacts in cloned sequences (Speksnijder et al., 2001) and thus might cause (at least a part of) the high degree of microheterogeneity observed for AOB sequence clusters detected in environmental 16S rRNA and amoA clone libraries. Although the 16S rRNA and amoA library approaches provided valuable insights into the natural diversity of AOB, the dependency of these techniques on nucleic acid extraction, PCR and cloning heavily biases the results (Wintzigerode and Goebel, 1997). Understand-

794

H.-P. Koops et al.

CHAPTER 3.2.12

Table 4. Phylogenetic affiliation of amoA sequences obtained from environmental samples. AmoA-sequences analyzed for this Table were obtained from Baribeau et al. (2000), Chang et al. (2001), Dionisi et al. (2002), Gieseke et al. (2001), Ivanova et al. (2000), Helmer et al. (1999), Henckel et al. (1999), Holmes et al. (1999), Horz et al. (2000), Nold et al. (2000), Oved et al. (2001), Purkhold et al. (2000), Purkhold et al. (2003), Rotthauwe et al. (1997), Sakano et al. (2002), Schmid et al. (2000), and Stephen et al. (1999). In addition, unpublished environmental amoA sequences determined by the authors or available in GenBank (Accession numbers: AJ277459], AF338319, AF338320], AF239878, AF239879, AF239880, AF239881, AF239882, AF239883, AF239884], AJ388568, AJ388569, AJ388570, AJ388571, AJ388572, AJ388573, AJ388574, AJ388575, AJ388576, AJ388577, AJ388578, AJ388579, AJ388580, AJ388581, AJ388582, AJ388583, AJ388588, and AJ388589) were used.

Nitrosospira cluster N. marina and N. oligotropha cluster N. europaea/Nc. mobilis cluster N. communis cluster Nitrosomonas sp. Nm143 cluster N. cryotolerans cluster Σ

Total

Isolates

Enrichments

Environmental clones

152 116 156 37 1 1 463

37 16 16 6 1 1 77

3 0 0 0 0 0 3

112 100 140 31 0 0 383

Nitrosospira lineage (23) N. oligotropha + N. marina lineage (24)

N. communis lineage (9)

N. europaeal Nc.mobilis lineage (23)

0%

20%

40%

60%

80%

soil (including rice roots, n=13)

marine environment (n=2)

wastewater treatment plant (n=22)

estuary (n=2)

freshwater environment (n=7)

biofilter (n=1)

100%

Fig. 13. Environmental distribution patterns of AOB clusters based on AmoA analysis. For each cluster, the origin (sampling site) of each AmoA sequence belonging to this cluster was determined. This analysis only included sequences from enrichments and sequences directly retrieved from the environment. Subsequently, we counted the number of different sampling sites in which members of a particular cluster were detected. This number, which is given in parentheses after the cluster designation, was set to 100%. In the next step, the sampling sites were grouped into the categories wastewater treatment plants, soil, freshwater (including sediment), estuary, biofilter, and marine (including sediment). For each cluster, the proportion of each category is shown. The displayed legend explains the color-coding of the habitats. In addition, the number of different sampling sites of a particular habitat category analyzed by the 16S rRNA approach (n) is given. Please note that the depicted grouping does not reflect the phylogenetic relation but results from the poor resolution of the amplified amoA gene fragment.

ing the ecology of AOB requires quantitative data on their in situ abundance and activity in ecosystems. The AOB community composition in a system can be quantitatively analyzed by using rRNA-targeted probes in combination with dot blot or in situ hybridization techniques (see the section on Molecular Techniques in this Chapter). However up to now, the AOB composition of only wastewater treatment reactors and full-scale plants has been characterized with these methods (Wagner et al., 1995; Mobarry et al., 1996; Juretschko et al., 1998;

Schramm et al., 1999; Schramm et al., 2000; Daims et al., 2001b).

Methods Useful for In Situ Detection of AOB Classical Techniques Traditionally, the abundance, diversity and ecology of AOB were investigated by cultivationdependent methods. For the enumeration of

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

viable AOB, the most probable number (MPN) technique often has been used, sometimes in parallel with other techniques. The efficiency of MPN has been calculated from comparisons with results obtained with fluorescent antibody (FA) counting (Belser, 1979; Ward, 1982) or calculations of activity per cell (Belser and Mays, 1982). Results obtained from these comparisons have led to the suggestion that the MPN efficiency may range between 0.001 and 0.05% (Hall, 1986). One important reason of underestimating AOB with MPN may be that AOB often form cell aggregates in the environment (Wagner et al., 1995; Stehr et al., 1995b; Juretschko et al., 1998; Koops and Pommerening-Röser, 2001). Another reason may be that the media used in MPN counting of AOB often have not been well adapted (ammonia concentration, salinity, and pH value) to the conditions in the environment under investigation (Koops and PommereningRöser, 2001). Qualitative characterization of natural AOB populations is a very time consuming process. This especially holds true if classical methods are applied. The isolation of numerically dominant species will take at least some months (see the section on Enrichment, Isolation, and Maintenance in this Chapter). Moreover, considerable experience in working with AOB in the laboratory is required. However, this approach is currently the only option that allows the physiological properties of those species representing the numerically dominant AOB population in the respective environment to be completely investigated. These findings can help determine why distinct species of AOB are dominant in certain environments. Phenotypic identification of new isolates of AOB is possible if the strain belongs to one of those species already being described in the literature. Useful parameters are the physiological properties (Ks values of ammonia-oxidation, requirement or tolerance of increased salt concentrations, and possession of urease), morphological characteristics (shape and size of cells, arrangement of intracytoplasmic membranes, and possession of carboxysomes), and G+C content of the DNA. For unambiguous identification, DNA-DNA hybridization with the type strain of the respective species should be carried out.

Immunological Techniques In the 1970s, the cultivation-dependent approach was complemented by the fluorescent antibody technique, which allowed (for the first time) members of this guild to be detected in situ. Generally, polyclonal antibodies against several AOB

795

were produced by injecting rabbits with the respective cells. Several studies demonstrated in situ visualization of AOB in environmental samples using these antisera for immunofluorescence and claimed that this approach allows a more rapid and accurate enumeration of AOB compared to enrichment culture methods (Belser, 1979; Ward and Perry, 1980; Ward and Carlucci, 1985; Abeliovich, 1987; Völsch et al., 1990). Although the fluorescent antibody (FA) approach represented a significant step forward in the history of research on nitrifier ecology, this technique has several disadvantages. Most importantly, pure cultures of AOB from the environment under investigation are required to produce the antibodies, since different serological groups exist within at least some of the AOB species. Therefore, the FA-technique, although allowing direct in situ detection of AOB, is indirectly dependent on successful cultivation of the target organisms. Consequently, this technique will not allow identification of novel, previously uncultured AOB species. Furthermore, the various polyclonal antibodies raised in different laboratories against different AOB are not easily available and thus their use is often restricted to a few laboratories. Another major disadvantage of the FA-technique is that the specificity of the obtained antibodies cannot be tailored in silico (i.e., using a computer) and must be determined in time-consuming experimental screening assays using different pure cultures. Finally, the FA-technique potentially suffers from false negative and false positive signals. The false negative signals might result from penetration problems of the relatively large antibodies in dense flocs or biofilms. False-positive signals were described as a result from nonspecific antibody binding to extracellular material or detritus (Szwerinski et al., 1985). Recently, polyclonal antibodies against the purified β subunit of the ammonia monooxygenase (AmoB) of Nitrosomonas eutropha were shown to target all betaproteobacterial AOB by Western blot analysis (Pinck et al., 2001). After appropriate AOB cell permeabilization (Bartosch et al., 1999), the α AmoB antibodies are also suitable for immunofluorescence-based group-specific detection of betaproteobacterial AOB (E. Spieck, personal communication). Since these polyclonal antibodies target a key enzyme of the betaproteobacterial AOB, this assay is expected to be highly specific and will probably also allow yet unknown members of this group to be targeted. In the future, this technique most probably can be standardized to become the first choice method for general counting of betaproteobacterial AOB in those environmental samples that are well suited for FA application.

796

H.-P. Koops et al.

Molecular Techniques After the establishment of 16S rRNA-based phylogeny of cultured AOB (Head et al., 1993; Teske et al., 1994; Pommerening-Röser et al., 1996; see the section on Phylogeny of AOB in this Chapter), many different 16S rRNA-targeted PCR primers and probes for dot blot or fluorescence in situ hybridization (FISH) have been developed (Table 5). Most frequently the natural diversity of AOB has been investigated by comparative analyses of PCR amplified and cloned AOB 16S rRNA gene fragments (see the section on Distribution of AOB in Nature in this Chapter). Apparently, the specificity of the applied PCR primers dramatically determines the outcome of such molecular AOB diversity surveys. In this context, it is important to note that the actual specificity of many primers used in previous investigations significantly differs from the indicated (intended) specificity listed in the original publications (Tables 6 and 7). These inconsistencies were detected after re-evaluating the primer/probe specificities based on the current, encompassing AOB 16S rRNA gene database. For example the PCR primer NitA, which is frequently used for AOB diversity surveys (Abd El Haleem et al., 2000; Ward et al., 2000; Hollibaugh et al., 2002), shows mismatches to all cultured nitrosospiras, all members of the N. communis and N. oligotropha lineage, and most members of the N. marina lineage. To avoid such pronounced biases, the use of PCR primers with high sensitivity (targeting all known or at least most betaproteobacterial AOB) but relatively low specificity (not perfectly excluding other bacteria) is recommended. Furthermore, the amplified 16S rRNA gene fragment must have a size larger than 1000 nucleotides to allow reliable phylogeny inference. The above-mentioned criteria are currently best fulfilled by the β AMOf and β AMOr primer pair. This approach accepts the amplification of non-AOB 16S rRNA gene fragments, which subsequently have to be identified by phylogenetic analysis or hybridization with probes with excellent specificity (e.g., probe Nso1225). Recently, the toolbox for AOB detection has been extended by analyzing PCR amplified 16S rRNA gene fragments with denaturing gradient gel electrophoresis (DGGE; Kowalchuk et al., 1997; Nicolaisen and Ramsing, 2002). This fingerprinting approach allows a rapid comparative assessment of AOB diversity in many samples but requires subsequent controls like cloning and sequencing of bands or hybridization of membrane-blotted DGGE gels with suitable probes. Furthermore, the length of PCR amplificates that can be analyzed is limited to approximately 500 bp, which limits primer selection and hampers phylogeny inference.

CHAPTER 3.2.12

Quantitative analysis of AOB community composition and abundance in the environment requires the use of methods not affected by PCR biases (Wintzigerode and Goebel, 1997). For this purpose, both dot blot and FISH methods have been developed and applied for AOB diversity research (Wagner et al., 1995; Mobarry et al., 1996; Schramm et al., 1996; Juretschko et al., 1998; Okabe et al., 1999; Liebig et al., 2001; Nogueira et al., 2002). The major advantage of dot blot analysis is that it can be applied to almost all environmental samples, while FISH does not work quantitatively in samples having a high autofluorescence or containing nontransparent particles (like soils). On the other hand, FISH allows visualization of the target bacteria and quantification of the AOB community composition and absolute AOB cell numbers (Daims et al., 2001c), while dot blot analysis only provides information on the relative or absolute abundance of probe target rRNA molecules. Whether, for example, a high rRNA content of a certain AOB species in an environmental sample originates from many target cells with a low rRNA concentration or a few cells with a high rRNA concentration cannot be figured out by dot blot hybridization. It has been postulated that the rRNA content of a bacterial cell is directly related to its physiological activity. It must however be noted that this link does not always hold true for AOB. Quantitative FISH experiments showed that AOB have high cellular ribosome contents even after prolonged complete physiological inhibition (Wagner et al., 1995) or starvation (Morgenroth et al., 2000). Therefore, physiological activity of AOB cannot reliably be inferred from the above-mentioned methods and must be determined for example by combining FISH and microautoradiography (Lee et al., 1999), FISH and microelectrodes (Schramm et al., 1999), or stable isotope probing (Whitby et al., 2001a). In addition to rRNA-based methods, the gene encoding the active site polypeptide of the ammonia monooxygenase (amoA) is frequently used as phylogenetic marker for AOB in environmental diversity surveys. Like the 16S rRNA database, the database of amoA sequences of cultured AOB is encompassing (see the section on Phylogeny of AOB in this Chapter). Several primer pairs for specific amplification of amoA have been published (Holmes et al., 1995; Rotthauwe et al., 1995; Sinigalliano et al., 1995; Mendum et al., 1999; Webster et al., 2002) but only the primer pair described by Rotthauwe et al. (1995), amplifying 55% of the amoA gene and slightly modified later by Stephen et al. (1999), has been shown to be suitable for detecting all tested betaproteobacterial AOB (Purkhold et al., 2000). The amoA approach for

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

797

Table 5. List of published 16S rRNA-targeted primers and probes. Probes that have been successfully used for FISH are shaded. Listed are specificities of primers and probes as given in the original publications. For an overview of the actual number of mismatches of each primer or probe with the 16S rRNA of all ammonia-oxidizer isolates please refer to Tables 6 and 7. Probe/primer

Target site

Specificitya

NM-75 NS-85 NmIIb NSMR32f NSMR71f NSMR34 NSMR76b NitA βAMOf NSPM Nm0 Nsm 156b NmVb CTO189f A/B-GC CTO189f C-GC Nso 190b Noli191b TAOfwd NM198 NmoCL6a_205 NmI Nmo218b TMP1 β-AO233 NspCL1_249 Nmo254a Nmo254 RT1r AAO258 Primer 356f NmoCL6b_376 Nsp436 NmoCL7_439 Nm439 NitD NMOB1f NSMR52f Nsv 443b NspCL4_446 Nsp0 NspCL3_454 NspCL2_458 Nlm 459r NSM1B Primer 517r TAOrev CTO654r NITROSO4Eb NEUb Amβ NitF NitC NmIII NSMR53r NSMR74r NMOB1r NSMR33r RNM-1007 NS-1009 NmIVb NitB Nso 1225b βAMOr Nse 1472b

67–86 76–95 120–139 125–141 126–143 131–149 132–149 136–158 142–162 145–162 148–165 155–173 174–191 189–207 189–207 189–207 191–208 192–208 198–218 205–221 210–225 218–236 226–253 233–249 249–266 254–271 254–271 283–304 258–277 356–372 376–392 436–453 439–456 439–459 439–461 442–461 443–461 443–461 446–463 452–469 454–471 458–475 458–477 478–494 517–533 632–649 632–653 638–657 651–668 738–758 844–862 846–862 998–1018 999–1017 1000–1017 1006–1026 1006–1021 1005–1028 1007–1026 1004–1022 1213–1233 1224–1243 1295–1314 1472–1489

Terrestrial Nitrosomonas spp./Nitrosococcus mobilis Nitrosospira spp. Nitrosomonas communis lineage Nitrosospira tenuis-like AOB Nitrosomonas marina-like AOB Nitrosospira tenuis-like AOB Nitrosomonas marina-like AOB βAOB βAOB βAOB Nitrosomonas spp. Nitrosomonas spp./Nitrosococcus mobilis Nitrosococcus mobilis βAOB βAOB βAOB Nitrosomonas cluster 6a Terrestrial AOB Nitrosomonas ureae + Nitrosomonas sp. AL212 Nitrosomonas cluster 6a Nitrosomonas europaea-lineage Nitrosomonas oligotropha-lineage AOB βAOB Nitrosospira cluster 1 All Nitrosomonas All Nitrosomonas AOB Terrestrial βAOB Nested PCR in NitAB amplicons Nitrosomonas cluster 6b All Nitrosospira Nitrosomonas cluster 7 Nitrosomonas ureae + Nitrosomonas sp. AL212 Nitrosomonas europaea Nitrosococcus mobilis-like AOB Nitrosomonas europaea-like AOB Nitrosospira spp. Nitrosospira cluster 4 Nitrosospira spp. Nitrosospira cluster 3 Nitrosospira cluster 2 Nitrosospira multiformis/Nitrosospira sp. C-141 Nitrosomonas europaea-lineage/Nitrosococcus mobilis Nested PCR in NitAB amplicons Terrestrial ammonia oxidizers βAOB βAOB Most halophilic and halotolerant Nitrosomonas βAOB βAOB βAOB Nitrosomonas marina-lineage Nitrosomonas europaea-like AOB Nitrosomonas marina-like AOB Nitrosococcus mobilis-like AOB Nitrosospira tenuis-like AOB Terrestrial Nitrosomonas spp. Nitrosospira spp. Nitrosomonas cryotolerans-lineage βAOB βAOB βAOB Nitrosomonas europaea-lineage

References Hiorns et al., 1995 Hiorns et al., 1995 Pommerening-Röser et al., 1996 Burrell et al., 2001 Burrell et al., 2001 Burrell et al., 2001 Burrell et al., 2001 Voytek and Ward, 1995 McCaig et al., 1994 Silyn-Roberts and Lewis, 2001 Pommerening-Röser et al., 1996 Mobarry et al., 1996 Pommerening-Röser et al., 1996 Kowalchuk et al., 1997 Kowalchuk et al., 1997 Mobarry et al., 1996 Gieseke et al., 2001; Rath, 1996 Chandler et al., 1997 Suwa et al., 1997 Stephen et al., 1998 Pommerening-Röser et al., 1996 Gieseke et al., 2001 Hermansson and Lindgren, 2001 Stephen et al., 1998 Stephen et al., 1998 Stephen et al., 1998 Stephen et al., 1998 Hermansson and Lindgren, 2001 Hiorns et al., 1995 Hollibaugh et al., 2002 Stephen et al., 1998 Stephen et al., 1998 Stephen et al., 1998 Suwa et al., 1997 Ward et al., 1997 Burrell et al., 2001 Burrell et al., 2001 Mobarry et al., 1996 Stephen et al., 1998 Pommerening-Röser et al., 1996 Stephen et al., 1998 Stephen et al., 1998 Hastings et al., 1997 Hovanec and DeLong, 1996 Hollibaugh et al., 2002 Chandler et al., 1997 Kowalchuk et al., 1997 Hovanec and DeLong, 1996 Wagner et al., 1995 Utaaker and Nes, 1998 Ward et al., 1997 Voytek and Ward, 1995 Pommerening-Röser et al., 1996 Burrell et al., 2001 Burrell et al., 2001 Burrell et al., 2001 Burrell et al., 2001 Hiorns et al., 1995 Hiorns et al., 1995 Pommerening-Röser et al., 1996 Voytek and Ward, 1995 Mobarry et al., 1996 McCaig et al., 1994 Juretschko et al., 1998

a Listed are specificities of primers and probes as given in the original publications. For an overview of the actual number of mismatches of each primer or probe with the 16S rRNA of all ammonia-oxidizer isolates, please refer to Tables 6 and 7. b Probe has been successfully used for FISH.

5 >10 0 0 0 0 3 0 >10 >10 5 2 2 7 3 2 >10 8 9 1 0 >10 >10 >10 >100 >100 >100 >100 >100 >100 4 0 1 0 0 0 2 >10 1 1 4 1 >10 >100 5 3 >10 3 >10 0 1 0 0 1 0 3 0 1 0 >10 4 >100 1

>10 6 0 0 0 0 0 0 7 0 1 2 0 2 0 2 3 0 0 0 0 5 >10 1 >100 >100 >10 >10 >100 >100 0 0 0 0 0 0 1 >10 1 0 0 1 6 >100 2 4 2 0 1 0 0 1 0 0 0 0 0 1 0 5 2 >10 0

NM-75 NS-85 NmII NSMR32f NSMR71f NSMR34 NSMR76 NitA bAMOf NSPM Nm0 Nsm 156 NmV CTO189f A/B-G CTO189f C-GC Nso 190 Noli191 TAOfwd NM198 NmoCL6a_205 NmI Nmo218 TMP1 b-AO233 NspCL1_249 Nmo254a Nmo254 AAO258 RT1r primer 356f NmoCL6b_376 Nsp436 NmoCL7_439 Nm439 NitD NMOB1f NSMR52f Nsv 443 NspCL4_446 Nsp0 NspCL3_454 NspCL2_458 Nlm 459r NSM1B primer 517r TAOrev CTO654r NITROSO4E NEU Amb NitF$ NitC$ NmIII NSMR53r NSMR74r NMOB1r NSMR33r RNM-1007 NS-1009 NmlV* NitB Nso 1225 bAMOr Nse 1472

1MM

0MM

Primer/Probe

NTO NTO

Cluster 2

2

Nsp Nsp AHB III2 40KI D11 GM4 III7 B6 T7 O13 O4 12 5 1

Cluster 0 Nsp 1

Nsp 4

GS 833 F3 E12

TYM 9

NRS 527

Nsp 40

Nsp 10 Nsp 62

1 1

Np 22

Cluster 3

3

6 4

6 4

NL 13

1 1

1

PJ TCH A1 716

1 1

1

1 1 1 1 1

RY 6A

1 1

1

2

1

RY 3C Ka3

Ka4

Cluster 4 Nsp Nsp 57 58

Nsp 65

H.-P. Koops et al.

3

2

2

1

1

1 1 1

Np Nv Nv L C Nsp Nsp Nsp A4 AF A16 24C NL5 Nv1 3919 6 12 115 141 AV 2 17

most sequences in cluster 1 are targeted

Nsp 41

Nitrosospira Isolates

Table 6. Specificity and sensitivity of published 16S rRNA (gene) targeting PCR primers and hybridization probes for nitrosospiras. Abbreviations: NTO, number of non-target organisms outside the AOB with no mismatch (0 MM) or one mismatch (1 MM) in its 16S rRNA (gene) to the respective primer/probe.a

798 CHAPTER 3.2.12

5 >10 0 0 0 0 3 0 >10 >10 5 2 2 7 3 2 >10

8 9 1 0 >10 >10 >10 >100 >100 >100 >100 >100 >100 4 0 1 0 0 0 2 >10 1 1 4 1 >10 >100 5 3 >10 3 >10 0 1 0 0 1 0 3 0 1 0 >10 4 >100 1

>10 6 0 0 0 0 0 0 7 0 1 2 0 2 0 2 3

0 0 0 0 5 >10 1 >100 >100 >10 >10 >100 >100 0 0 0 0 0 0 1 >10 1 0 0 1 6 >100 2 4 2 0 1 0 0 1 0 0 0 0 0 1 0 5 2 >10 0

NM-75 NS-85 NmII NSMR32f NSMR71f NSMR34 NSMR76 NitA bAMOf NSPM Nm0 Nsm 156 NmV CTO189f A/B-GC CTO189f C-GC Nso 190 Noli191 TAOfwd NM198 NmoCL6a_205 NmI Nmo218 TMP1 b-AO233 NspCL1_249 Nmo254a Nmo254 AAO258 RT1r primer 356f NmoCL6b_376 Nsp436 NmoCL7_439 Nm439 NitD NMOB1f NSMR52f Nsv 443 NspCL4_446 Nsp0 NspCL3_454 NspCL2_458 Nlm 459r NSM1B primer 517r TAOrev CTO654r NITROSO4E NEU Amb NitF$ NitC$ NmIII NSMR53r NSMR74r NMOB1r NSMR33r RNM-1007 NS-1009 NmlV* NitB Nso 1225 bAMOr Nse 1472

1MM

0MM

Primer/Probe

NTO NTO

WH 2

Nm ENI- Nm 11 103 50

Koll 21

Nm 57

GH 22

CNS G1 K1 326

IWT 202

IWT 514

1

DYS 317

1

DYS 323

TK F6 F5 794

N. europaea /Nc. mobilis Cluster HPC 101

AN 1-5 Nm 1

Nc 2

Nm Nm 93 104

Nm 107

TNO 632

1

Nm 90

Nm Nm 148 2

Nm 33

Nm 41

Nm 58

N. communis Cluster TA 921

3

B19

Nitrosomonas Isolates Nm 51

Nm 63 Nm 22

2 2 2 1 1

C 113

C 45

1

2

1

mar Nm NH4W URW NO3W 122 36

N. marina Cluster Nm 47

2

Nm 45

Nm 10

1

AL Nm 212 80

JL 21

Nm 59

N. oligotropha Cluster Nm 84

1

TT140 TT140 89A 098-2

Nm 143

Nm143 Cluster C17

Table 7. Specificity and sensitivity of published 16S rRNA (gene) targeting PCR primers and hybridization probes for nitrosomonads. Abbreviations: NTO, number of non-target organisms outside the AOB with no mismatch (0 MM) or one mismatch (1 MM) in its 16S rRNA (gene) to the respective primer/probe.a

Nm 55

CHAPTER 3.2.12 The Lithoautotrophic Ammonia-Oxidizing Bacteria 799

800

H.-P. Koops et al.

AOB diversity research impresses by its specificity and sensitivity but suffers from PCR-biases and relatively low phylogenetic information content (see the section on Phylogeny of AOB in this Chapter). Several fingerprinting techniques like gel retardation, DGGE, and terminal restriction fragment length polymorphism (T-RFLP) have been combined with the amoA approach and allow first insights into the diversity of an amoA PCR amplificate without time-consuming cloning and sequencing analysis (Kowalchuk et al., 1998; Horz et al., 2000; Schmid et al., 2000; Nicolaisen and Ramsing, 2002). Furthermore, an in situ PCR technique is available which allows direct detection of the amoA gene in AOB (Hoshino et al., 2001). Significant method development for AOB diversity research is underway. Several quantitative PCR methods for AOB detection in the environment have been developed (Mendum et al., 1999; Dionisi et al., 2002). Furthermore, absolute AOB numbers in complex samples can also be determined by applying a novel FISH method in combination with confocal laser scanning microscopy and digital image analysis (Daims et al., 2001c). In the future, DNA microarrays composed of several hundred rRNA- or amoA -targeted oligonucleotide probes will almost certainly become available for parallel analyses of AOB community structures. Studies on first generation microarrays carrying a relatively limited number of AOB-targeted probes have already been published (Guschin et al., 1997; Wu et al., 2001).

Distribution of AOB in Nature The distribution of AOB in the environment has traditionally been analyzed by cultivationdependent methods (see the section on Classical Techniques in this Chapter). Recently, these studies were complemented by molecular diversity surveys of various man-made and natural systems (see the section on Molecular Techniques in this Chapter). The combination of both approaches is strongly recommended because each approach is characterized by its specific limitations. MPN-based isolation of AOB target the numerically most important AOB but might overlook not yet cultured AOB species that cannot thrive in the cultivation medium offered (although any clear indication for that is missing in the literature). On the other hand, it is important to interpret PCR-based molecular AOB distribution data with caution because the mere retrieval of a 16S rRNA sequence of an AOB proves neither that this organism is abundant nor that it is physiologically active. This problem is particularly severe for AOB (com-

CHAPTER 3.2.12

pared to many other bacteria) because they can survive very long time periods under unfavorable conditions (Johnstone and Jones, 1988; Wilhelm et al., 1998; Pinck et al., 2001). Therefore, molecular studies aiming at identification of dominant AOB species in an environment should focus on PCR amplificates of the highest positive DNA dilution, an approach analogous to the MPN technique (Feray et al., 1999), or even better include quantitative techniques (see the section on Molecular Techniques in this Chapter). Independent of the method used, the distinction between autochthonous and allochthonous AOB of a particular ecosystem is not trivial, making the natural distribution patterns of the distinct AOB species difficult to define. This problem is particularly severe because many of the environments are strongly influenced by human activity that eventually causes transmission of significant amounts of AOB to foreign systems. For example, AOB typically found in activated sludge also might be detectable in freshwater systems, if effluents of a wastewater treatment plant are disposed close to the sampling site. In this context, the importance of a careful selection of representative sampling sites is obvious. It is also interesting to note that isolations of strains via MPN series in general define more restricted distribution patterns of the distinct species of AOB than obtained from comparable molecular investigations. AOB are present in most aerobic environments including rocks and building stones but can also survive in anaerobic systems (see the section on Dominant Populations of AOB in Different Environments in this Chapter). Since ammonia is the essential energy source for these organisms, their distribution in nature is coupled to geological, biological and anthropogenic sources of reduced nitrogen. Consequently, they have adapted to a broad range of different ammonia concentrations in the diverse environments, reflected by different affinity constants for ammonia. This is one of the most important aspects influencing the distribution patterns of AOB in nature (Suwa et al., 1994; Suwa et al., 1997; Stehr et al., 1995a; Koops and Pommerening-Röser, 2001). On the other side, ammonia is a toxic compound. Therefore, the tolerance of increasing ammonia concentrations (Table 3) is another aspect shaping the distribution patterns of AOB (Bollmann and Laanbroek, 2001). Furthermore, the presence or absence of urease activity was observed to be of ecophysiological relevance for AOB. This property is of special importance in acidic environments, where free ammonia is missing as substrate because it is nearly quantitatively ionized to ammonium. Under such conditions, only those AOB species that can use urea as an alternative

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

ammonia source can build up stable populations (De Boer and Laanbroek, 1989; De Boer et al., 1991; Jiang and Bakken, 1999; Burton and Prosser, 2001). Possession of urease seems also to be an essential property for AOB that successfully colonize oligotrophic soils or aquatic environments. Distribution of AOB in nature also is affected by different salt requirements, salt tolerances and salt sensitivities of the distinct species (Table 3). This is of special importance for their distribution patterns in aquatic systems (such as rivers, lakes, estuaries, marine environments and salt lakes) that significantly differ in salinity (Koops et al., 1990; Koops et al., 1991). The AOB are aerobes. Although their oxygen affinity constants are relatively high (Painter, 1986), AOB possess nitrifying activity not only at oxygen saturation but also at extremely low oxygen concentrations (Goreau et al., 1980). Even ammonia oxidation under anaerobic conditions is being discussed (Schmidt and Bock, 1997; Schmidt and Bock, 1998; Zart et al., 2000). However, specific selection of distinct AOB species by different oxygen concentrations has not yet been reported in the literature. Temperature also may be of importance for distribution patterns of AOB in nature. This has been indicated in some publications (Golovacheva, 1976; Jones et al., 1988; Jiang and Bakken, 1999). Beside environments characterized by constant high or low temperatures, such as hot springs or permafrost soils, environments showing pronounced temperature changes, such as rock surfaces, might harbor interesting AOB. However, these systems have not been studied intensively enough to allow general conclusions on their AOB diversity.

The Betaproteobacterial AOB The betaproteobacterial AOB are divided into two main subgroups, the Nitrosomonas group (containing six distinct lineages) and the Nitrosospira lineage (containing the genera Nitrosospira, Nitrosovibrio and Nitrosolobus). Within the Nitrosomonas group, a relatively strong correlation between the phylogenetically defined lineages (see the section on Phylogeny of AOB in this Chapter) and the distribution patterns of the respective species exists. Within the Nitrosospira lineage, this correlation is not as obvious. The NITROSOMONAS Group The N. europaea/Nc. mobilis Lineage The four cultured species of the N. europaea/Nc. mobilis lineage are moderately halophilic and all show

801

striking halotolerance (Table 3). Most of the cultured strains of the three species N. europaea, N. eutropha and Nitrosococcus mobilis (which is phylogenetically a member of the genus Nitrosomonas) have been isolated from wastewater treatment plants and their molecular signatures are also frequently retrieved from these systems (Figs. 12 and 13). In addition, quantitative FISH results demonstrated that Nc. mobilis and N. europaea are dominant in several wastewater treatment plants (Juretschko et al., 1998; Daims et al., 2001b). Some isolates of this lineage were obtained from strongly fertilized agricultural soils. All these environments are more or less eutrophicated and influenced by anthropogenic activity, frequently rendering the conditions more favorable for these AOB. The successful colonization of these systems requires tolerance of increasing ionic strength and, in particular, of high concentrations of toxic compounds such as ammonia. Consequently, these properties are found in all members of these AOB lineages (Table 3). The natural habitat for members of these lineages has not yet been identified. However, in a recent molecular investigation the presence of N. europaea -like and N. eutropha -like organisms in a salt lake was shown (Ward et al., 2000). This is of interest, since strains belonging to the fourth species of the N. europaea/Nc. mobilis lineage, N. halophila, recently could be isolated from salt lakes in Mongolia (Sorokin et al., 2001). Sporadically, isolates of the N. europaea/Nc. mobilis lineage were obtained from the North Sea (N. halophila and N. mobilis), from the river Elbe estuary (N. europaea) and from freshwater sediments (N. europaea and N. eutropha ; Stehr et al., 1995a; Koops and Pommerening-Röser, 2001). Consistent with these findings, molecular investigations detected members of the N. europaea/Nc. mobilis lineage in fertilized soils or natural fresh water environments (Voytek, 1996; Holmes et al., 1999; Campbell, 2000; Horz et al., 2000; Ivanova et al., 2000; Phillips et al., 2000; Burrell et al., 2001; Oved et al., 2001; Smith et al., 2001; Whitby et al., 2001a; Morris et al., 2002; Radajewski et al., 2002; Webster et al., 2002). The N. communis Lineage The N. communis lineage comprises two entities, the N. communis and the N. nitrosa sublineages, which are currently phylogenetically not clearly distinguishable but have significantly different distribution patterns in nature. The N. communis sublineage contains, in addition to N. communis, three other cultivated species, not yet described by name (Fig. 9). The majority of strains of this sublineage were isolated from approximately pH neutral soils, often agriculturally utilized. Since all three species of this sublineage possess relatively high

802

H.-P. Koops et al.

affinity constants for ammonia and lack urease activity, they probably need relatively high ammonia concentrations in the environment. Most likely this is an important reason that these species seem to be missing in oligotrophic natural soils and in substrate-low natural freshwaters. In contrast to the above subgroup, strains of the N. nitrosa subgroup are commonly distributed in more or less eutrophicated freshwaters. Since the affinity constants for ammonia are similar to those of the species of the N. communis sublineage, the possession of urease might be responsible for this different distribution. Strains of N. nitrosa also have been isolated from marine environments, a fact which is in apparent contrast to the relatively low salt tolerance of these strains observed in laboratory studies (Table 3). Surprisingly, only two 16S rRNA gene clones of the N. communis lineage have been detected in environmental diversity surveys (Gieseke et al., 2001; Fig. 11). This low retrieval rate may have been caused by the fact that many of the commonly applied primers possess mismatches with the 16S rRNA gene of members of this lineage (Table 7). In contrast, amoA sequences affiliated with this lineage were detected in soil, freshwater and wastewater treatment plants (Ivanova et al., 2000; Purkhold et al., 2000; Gieseke et al., 2001; Oved et al., 2001; Dionisi et al., 2002; Nogueira et al., 2002; Fig. 11). The N. oligotropha Lineage The N. oligotropha lineage comprises several species, although only N. oligotropha and N. ureae are described by name. A total of six species of this lineage is available in culture (Fig. 7), and the existence of more species has been indicated by molecular environmental studies (Speksnijder et al., 1998) and several unpublished sequences publicly accessible via GenBank. The majority of cultured strains have been isolated from rivers and lakes (Koops and Pommerening-Röser, 2001). Some isolates originate from sewage disposal plants (Koops and Harms, 1985; Suwa et al., 1997) and from soil samples. The 16S rRNA sequences of this cluster were frequently detected in soil and freshwater (Stephen et al., 1996; Speksnijder et al., 1998; Bruns et al., 1999; Radeva et al., 1999; Kowalchuk et al., 2000b; Phillips et al., 2000; Bollmann and Laanbroek, 2001; Whitby et al., 2001b; Regan et al., 2002). Among the cultivated species of AOB, the representatives of the N. oligotropha lineage reveal the lowest affinity constants for ammonia, being in the range of a few µM. All investigated strains are urease positive. These observations are in accordance with the generally low ammonia concentration in their natural habitats. It is also notable, that representatives of this lineage often occur floc- or biofilm-attached in the environ-

CHAPTER 3.2.12

ment (Stehr et al., 1995a), as well as in laboratory studies (Bollmann and Laanbroek, 2001). Consistent with this observation, at least some members of this lineage can produce remarkable amounts of exopolymeric substances (Stehr et al., 1995b). The N. marina Lineage The N. marina lineage comprises three cultivated species, N. marina, N. aestuarii and Nitrosomonas sp. (represented by the isolates Nm 51 and Nm 63). All isolates originate from marine environments. Molecular signatures of this lineage have also been detected in marine samples (Stephen et al., 1996; Casciotti and Ward, 2001) as well as in enrichment cultures from such environments (Stephen et al., 1996). The presence of members of this lineage has been observed with molecular techniques in estuarine environments, coastal sand dunes and even in freshwater (aquaria) and a wastewater treatment plant (Kowalchuk et al., 1997; Speksnijder et al., 1998; Purkhold et al., 2000; Burrell et al., 2001; de Bie et al., 2001; Nicolaisen and Ramsing, 2002). Some authors, unfortunately, have lumped together the N. oligotropha lineage and the N. marina lineage (Kowalchuk and Stephen, 2001), although both lineages are phylogenetically distinct (Figs. 6 and 7) and the species of the two lineages show pronounced differences in ecophysiological characteristics and distribution patterns. The N. cryotolerans Lineage The N. cryotolerans lineage is represented by only one species, N. cryotolerans, and only one isolate exists in culture. This strain has been isolated from a marine environment and is obligatorily halophilic and urease positive. In molecular in situ analyses of AOB populations, N. cryotolerans has not yet been detected, although most of the commonly applied 16S rRNA gene primers do match this species. The Nm 143 Lineage Recently, a new lineage was identified within the genus Nitrosomonas (Purkhold et al., 2003; Fig. 7). This lineage has been provisionally named after Nitrosomonas sp. Nm143 and contains three additional isolates. It is not known whether these four isolates represent one or more than one species. All isolates were obtained from marine systems. Molecular signatures of this lineage were found in several marine and estuary systems (McCaig et al., 1994; Stephen et al., 1996; de Bie et al., 2001; Nicolaisen and Ramsing, 2002). Uncultured Lineage The so-called “environmental lineage 5” is the only major evolutionary lineage within the Nitrosomonas group which does not yet contain a cultured representative

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

(Stephen et al., 1996; Purkhold et al., 2000). The 16S rRNA sequences of this lineage were obtained from freshwater environments, marine systems and a coastal sand dune (McCaig et al., 1994; McCaig et al., 1999; Stephen et al., 1996; Kowalchuk et al., 1997; Speksnijder et al., 1998; Bano and Hollibaugh, 2000).

803

At least within the so-called “Nitrosospira cluster 3,” profound differences regarding physiology and distribution patterns exist because this cluster, in contrast to the other four clusters, comprises many morphologically as well ecophysiologically different species.

The Gammaproteobacterial AOB The Nitrosospira Lineage Members of the Nitrosospira lineage were postulated by some authors to be the dominant AOB in nature (Hiorns et al., 1995; Hastings et al., 1997; Ceccherini et al., 1998). However, this postulate is rather unsecured because it is based on findings obtained with nonquantitative PCRdependent methods often involving primers and probes not sufficiently covering some of the Nitrosomonas lineages (Tables 6 and 7). The Nitrosospira lineage represents a phylogenetically young group of AOB. Therefore, no highly reproducible phylogenetic substructure can be revealed within this group based on comparative 16S rRNA or amoA sequence analysis (see the section on Phylogeny of AOB in this Chapter). Nevertheless, a provisional subdivision system of this group has been suggested (Stephen et al., 1996; extended by Purkhold et al., 2000; Fig 8) and widely applied for interpretation of data from molecular AOB diversity surveys (for a review, see Kowalchuk and Stephen, 2001). However, it is questionable whether the suggested clustering defines units characterized by particular ecophysiological traits. Most isolates of the Nitrosospira lineage were obtained from terrestrial systems. Consistent with this finding, the current molecular dataset suggests that members of the Nitrosospira clusters 0, 2, 3 and 4 are predominantly found in terrestrial habitats (Fig. 12). Whether the reported occurrence of these clusters in freshwater habitats reflects that these bacteria are autochthonous freshwater inhabitants or soilborne organisms transferred to the water bodies cannot be decided. Nitrosospiras of cluster 1, for which no isolate exists, also occur in soil but are more frequently found in marine systems. The analyzed wastewater treatment plants rarely harbored nitrosospiras, and only members of clusters 2 and 3 were detected. Constructed wetlands might represent an exception to this rule, since mainly Nitrosospira -related 16S rRNA sequences were retrieved from a plant using rhizoremediation technology (Abd El Haleem et al., 2000). Nitrosospira strains Nsp57, 58 and 65, which were isolated from masonry and form two novel branches within this group (Fig. 8), are the only Nitrosospira isolates for which yet no closely related sequences were directly obtained from an environment.

The cultured strains of the two species of the genus Nitrosococcus, representing the gammaproteobacterial AOB, have been isolated from marine environments (all strains of N. oceani) and from a salt lagoon or from a salt lake (two isolates of N. halophilus), respectively. This habitat range is in accordance with the salt requirements and with the salt tolerances of the two species in laboratory experiments. Using immunological methods and molecular techniques, N. oceani was detected exclusively in marine environments but shown to be widely distributed (Ward and Perry, 1980; Ward, 1982; Ward and O’Mullan, 2002). Nitrosococcus halophilus has not yet been observed in nature via molecular techniques.

Dominant Populations of AOB in Different Environments Attempts to describe the dominant AOB populations of an environment should always take into careful consideration whether the retrieved species are autochthonous or allochthonous members of the analyzed ecosystem. Often species are introduced into environments from adjacent habitats. Since ecophysiological characteristics of distinct species of AOB often differ only minimally, introduced populations have a good chance to survive in the novel environment. Moreover, anthropogenic influence has created many new environments colonized by AOB that had no or only little time to adapt evolutionarily to these systems. In such cases, the sum of ecophysiological parameters will determine finally which of the species competing with each other will become the dominant representative.

Marine Environments A relatively clear situation is found in marine environments. In these ecosystems, generally those AOB are dominant which have a compatible salt requirement and a suitable salt tolerance. Consequently, the gammaproteobacterial AOB, N. oceani together with representatives of the Nitrosomonas marina lineage are the only AOB isolated with high frequency from marine environments. The second species of Nitrosococ-

804

H.-P. Koops et al.

cus, N. halophilus, seems to prefer environments with higher salt concentrations (salt lagoons or even salt lakes). Most of the above-mentioned organisms (except N. halophilus) can also be detected in marine systems using molecular methods (Stephen et al., 1996; Ward and O’Mullan, 2002). It is not clear why only one strain of N. cryotolerans, which also is ecophysiologically well adapted to these systems, has been isolated from marine samples. Members of the Nm143 lineage also seem to be distributed exclusively in marine environments. However, the ecophysiological properties of the latter group have not yet been investigated. Occasionally, species of the Nitrosomons europaea/Nc. mobilis lineage, owing to their relatively pronounced salt tolerance, were cultured from marine samples (Koops et al., 1976; Koops and Pommerening-Röser, 2001). Using quantitative dot blot, Hovanec and DeLong (1996) found that N. europaea -like bacteria were a major component of the microbial community in filters from seawater aquaria, but the primer (NSM1B) used for this analysis also has a full match to several members of the N. marina lineage (Tables 6 and 7). The 16S rRNA sequences of the Nitrosomonas environmental lineage 5 and Nitrosospira cluster 1, which both contain no cultured representatives, were frequently detected in marine environments (McCaig et al., 1994; McCaig et al., 1999; Stephen et al., 1996; Phillips et al., 1999; Bano and Hollibaugh, 2000; Hollibaugh et al., 2002; Fig. 12), but quantitative molecular data are missing. The retrieval of nitrosospiras in systems with elevated salt concentrations is surprising, since all cultured species of this lineage have a low salt tolerance in laboratory studies. The mere detection of 16S rRNA sequences of Nitrosospira cluster 1 in a marine environment does not prove that these organisms indeed possess a high salt tolerance and thrive in these systems. Alternatively, these nitrosospiras might be allochthonous inhabitants able to survive for extended periods in the ocean without significant physiological activity. It is thus important 1) to isolate and study a strain of this cluster and/or 2) to prove physiological activity of members of this cluster in situ for example by combination of FISH and microautoradiography (Lee et al., 1999).

Salt Lakes In salt lakes, the pH as well as the composition and concentration of salts vary significantly (Ward et al., 2000; Sorokin et al., 2001) thus creating different niches for AOB. Accordingly, different AOB were found in different salt lakes. From a salt lake in Saudi Arabia, a strain of

CHAPTER 3.2.12

Nitrosococcus halophilus was obtained (Koops et al., 1990), and from soda lakes located in Mongolia, alkali tolerant representatives of Nitrosomonas halophila have been isolated (Sorokin et al., 2001). In molecular analyses, N. europaea -like and N. eutropha -like organisms were detected in the hypersaline Mono Lake in California (Ward et al., 2000). However, the PCR primers used in this study (NitA and NitD) were not suited for a general AOB diversity study (Tables 6 and 7). To get a more general view of the AOB community composition in such environments, more investigations must be carried out.

Freshwater Environments In natural freshwater environments, members of the Nitrosomonas oligotropha lineage are generally the dominant AOB representatives. This is indicated by repeated isolation of members of this lineage from high MPN dilutions (Koops and Harms, 1985; Stehr et al., 1995a; Koops and Pommerening-Röser, 2001). Quantitative PCRindependent molecular data on AOB community composition in these systems are not available, but at least 16S rRNA sequences of this lineage were frequently detected in freshwater water bodies and sediments (Speksnijder et al., 1998; Radeva et al., 1999; Bollmann and Laanbroek, 2001; Whitby et al., 2001b; Regan et al., 2002) as well as in the Schelde estuary (Belgium and The Netherlands; de Bie et al., 2001). In addition, 16S rRNA sequences of Nitrosospira clusters 0, 2, 3 and 4 were harvested in molecular analyses of these habitats (Kowalchuk et al., 1998; Speksnijder et al., 1998; Whitby et al., 1999; Whitby et al., 2001b) and in a freshwater aquarium (Burrell et al., 2001). Moreover, using immunological methods, nitrosospiras were found in a eutrophic lake sediment (Smorzewski and Schmidt, 1991). However, these results do not necessarily indicate Nitrosospira dominance in freshwater systems because quantitative data are lacking, and several of the PCR primers applied in these studies do not sufficiently cover the N. oligotropha cluster (Purkhold et al., 2000; Tables 6 and 7). Occasionally, strains of N. europaea, N. eutropha and N. nitrosa have been isolated from eutrophicated rivers and eutrophic lakes (Koops and Pommerening-Röser, 2001). Furthermore, 16S rRNA sequences of N. europaea/Nc. mobilis, of Nitrosomonas environmental lineage 5 and of the N. marina lineage were retrieved from freshwater systems (Speksnijder et al., 1998; Whitby et al., 1999; Bollmann and Laanbroek, 2001; Burrell et al., 2001), but the significance of these findings remains unclear.

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

805

Artificial Aquatic Environments Urea and ammonia are the most frequently found nitrogen compounds in sewage. In wastewater treatment plants, AOB oxidize ammonia to nitrite, which is subsequently converted to nitrate by the nitrite-oxidizing bacteria. Nitrate is then removed from the sewage by denitrifying bacteria via anaerobic respiration. The slow growth rate of AOB and their susceptibility to pH- and temperature swings as well as to several sewage compounds is responsible for frequent failure of the nitrification in municipal and industrial wwtps. Isolation techniques indicate that in standard municipal wwtps, N. eutropha seems to be the dominant representative (Watson and Mandel, 1971b; Koops and Harms, 1985), but N. europaea and Nc. mobilis have also been repeatedly cultivated (Juretschko et al., 1998; Koops and Pommerening-Röser, 2001). All these species belong to the same Nitrosomonas lineage (Fig. 7) In industrial wwtps, the cultivation of representatives of members of the N. oligotropha lineage as well as N. nitrosa has been regularly reported (Koops and Harms, 1985; Suwa et al., 1997). In laboratory experiments, a remarkable high tolerance of members of the N. oligotropha lineage to heavy metals was observed, and the production of significant amounts of exopolymeric materials by these species was suggested to be the major reason for this tolerance (Stehr et al., 1995b). This resistance to heavy metals may be responsible for the presence of members of this lineage in special wwtps. During the last decade, molecular techniques were extensively applied to characterize AOB community structure and activity in activated sludge and biofilm from various wwtps. FISH revealed that AOB generally occur in tight microcolonies formed of up to several thousand individual cells (Fig. 14). These microcolonies cannot be easily dispersed using ultrasonic or Ultraturrax homogenizer treatments and thus contribute to the underestimation of AOB numbers in MPN assays. Typically, the AOB are detected in situ in close proximity to nitrite oxidizers reflecting the mutualistic relationship between both guilds (Mobarry et al., 1996; Schramm et al., 1996; Schramm et al., 1998; Juretschko et al., 1998; Okabe et al., 1999). Experiments using a combination of FISH with 14 C-bicarbonate microautoradiography demonstrated that almost all AOB microcolonies in activated sludge are physiologically active. Furthermore, AOB in wwtps can incorporate pyruvate (Daims et al., 2001a). Using quantitative FISH data, the specific activity of a single AOB cell in activated sludge has been estimated to be 2.3 ± 0.4 fmol of ammonia per hour (Daims

Fig. 14

Fig. 14. FISH detection of an AOB microcolony in a nitrifying biofilm from a sequencing batch biofilm reactor. AOB were stained by probe NEU, and the fluorescence signals were recorded by confocal laser scanning microscopy. Probeconferred fluorescence is shown in yellow, while autofluorescence of the biofilm is depicted in blue. Bar = 10 µm. The picture was kindly provided by Holger Daims.

et al., 2001c). According to FISH analyses, nitrosomonads are predominant over nitrosospiras in most activated sludge and biofilm systems (Mobarry et al., 1996; Schramm et al., 1996; Schramm et al., 2000; Juretschko et al., 1998; Okabe et al., 1999; Daims et al., 2001b; Gieseke et al., 2001). With the exception of Nitrosomonas cryotolerans, Nitrosomonas halophila and Nitrosomonas sp. Nm143, relatives of all other Nitrosomonas species occur in wwtps (Purkhold et al., 2000). Nitrosococcus mobilis, originally thought to be restricted to brackish water, is abundant in many wwtps (Juretschko et al., 1998; Daims et al., 2001b). Some nitrifying wwtps are dominated by a single AOB species (Juretschko et al., 1998), while other plants harbor a high diversity of AOB (Purkhold et al., 2000; Daims et al., 2001b; Gieseke et al., 2001). Whether and how differences in diversity of ammonia oxidizers are linked to the stability of the nitrification process is a yet unanswered question. Investigations of AOB populations in the raw sewage most probably also could give important insight into AOB diversity in sewage disposal plants.

Soils In acidic soils (heathlands, grasslands, forest soils, and marshy lands), where sufficient amounts of free ammonia are missing, ureasepositive species seem to be the exclusive representatives of the AOB. This has become obvious

806

H.-P. Koops et al.

via isolations as well as from molecular investigations (Kowalchuk and Stephen, 2001). Exclusively, species of the genera Nitrosospira and Nitrosovibrio could be detected. Molecular studies confirmed the predominance of nitrosospiras in acidic soils (Stephen et al., 1996; Smith et al., 2001) and indicated that, in particular, members of the Nitrosospira cluster 2 are well adapted to low pH terrestrial habitats (Kowalchuk et al., 1997; Stephen et al., 1998). A more detailed upto-date summary of these findings can be found in Kowalchuk and Stephen (2001). However, urease-positive strains of Nitrosolobus multiformis and of the genus Nitrosomonas, respectively, are missing in acidic soils. In more neutral soils, especially if agriculturally used, members of the N. communis lineage together with strains of Nitrosolobus multiformis are absolutely dominant in high MPN dilutions (MacDonald, 1986; Koops and PommereningRöser, 2001). In oligotrophic natural soils, especially if moderately acidic (pH around 6.0), strains of the N. oligotropha lineage as well as strains of the genera Nitrosospira or Nitrosovibrio have also been isolated (Koops and Pommerening-Röser, 2001).

Rocks and Building Stones AOB are also common on surfaces of rocks or building stones. Many strains have been isolated, and all were found to be members of the genera Nitrosospira or Nitrosovibrio (Spieck et al., 1992). Neither the closely related genus Nitrosolobus nor any of the lineages of the genus Nitrosomonas seem to be represented in these environments. Currently, the selective factors leading to these distribution patterns are speculative. Most likely, the acidic pH caused by the nitrification process is a crucial factor. Periodically appearing dryness or drastic change in temperature also may be selective for members of the genera Nitrosospira or Nitrosovibrio.

Literature Cited Aakra, A., J. B. Utaaker, I. F. Nes, and L. R. Bakken. 1999a. An evaluated improvement of the extinction dilution method for isolation of ammonia-oxidizing bacteria. J. Microbiol. Meth. 39:23–31. Aakra, A., J. B. Utaaker, and I. F. Nes. 1999b. RFLP of rRNA genes and sequencing of the 16S-23S rDNA intergenic spacer region of ammonia-oxidizing bacteria: A phylogenetic approach. Int. J. Syst. Bacteriol. 49(1):123–130. Aakra, A., M. Hesselsoe, and L. R. Bakken. 2000. Surface attachment of ammonia-oxidizing bacteria in soil. Microb. Ecol. 39:222–235. Aakra, A., J. B. Utaaker, and I. F. Nes. 2001a. Comparative phylogeny of the ammonia monooxygenase subunit A

CHAPTER 3.2.12 and 16S rRNA genes of ammonia-oxidizing bacteria. FEMS Microbiol. Lett. 205:237–242. Aakra, A., J. B. Utaaker, A. Pommerening-Roser, H.-P. Koops, and I. F. Nes. 2001b. Detailed phylogeny of ammonia-oxidizing bacteria determined by rDNA sequences and DNA homology values. Int. J. Syst. Evol. Microbiol. 51:2021–2030. Abd El Haleem, D., F. von Wintzingerode, A. Moter, H. Moawad, and U. B. Göbel. 2000. Phylogenetic analysis of rhizosphere associated β-subclass proteobacterial ammonia oxidizers in a municipal wastewater treatment plant based on rhizoremediation technology. Lett. Appl. Microbiol. 31:34–38. Abeliovich, A. 1987. Nitrifying bacteria in wastewater reservoirs. Appl. Environ. Microbiol. 53:754–760. Alzerreca, J. J., J. M. Norton, and M. G. Klotz. 1999. The amo operon in marine, ammonia–oxidizing gamma– proteobacteria. FEMS Microbiol. Lett. 180:21–29. Bano, N., and J. T. Hollibaugh. 2000. Diversity and distribution of DNA sequences with affinity to ammoniaoxidizing bacteria of the beta subdivision of the class Proteobacteria in the Arctic Ocean. Appl. Environ. Microbiol. 66:1960–1969. Baribeau, H., C. A. Kinner, J. R. Stephen, R. de Leon, P. A. Rochelle, and D. L. Clark. 2000. Presented at the American Water Works Association Water Quality Technology Conference Proceedings. Bartosch, S., I. Wolgast, E. Spieck, and E. Bock. 1999. Identification of nitrite-oxidizing bacteria with monoclonal antibodies recognizing the nitrite oxidoreductase. Appl. Environ. Microbiol. 65:4126–4133. Belser, L. W. 1979. Population ecology of nitrifying bacteria. Ann. Rev. Microbiol. 33:309–333. Belser, L. W., and E. L. Mays. 1982. Use of nitrifier activity measurements to estimate the efficiency of viable nitrifier counts in soilsand sediments. Appl. Environ. Microbiol. 43:945–948. Bollmann, A., and H. J. Laanbroek. 2001. Continuous culture enrichments of ammonia-oxidizing bacteria at low ammonium concentrations. FEMS Microbiol. Ecol. 37:211–221. Bruns, M. A., J. R. Stephen, G. A. Kowalchuk, J. I. Prosser, and E. A. Paul. 1999. Comparative diversity of ammonia oxidizer 16S rRNA gene sequences in native, tilled, and successional soils. Appl. Environ. Microbiol. 65:2994– 3000. Buchanan, R. E. 1917. Studies on the nomenclature and classification of bacteria. III: The families of the Eubacteriales. J. Bacteriol. 2:347–350. Burrell, P. C., C. M. Phalen, and T. A. Hovanec. 2001. Identification of bacteria responsible for ammonia oxidation in freshwater aquaria. Appl. Environ. Microbiol. 67:5791–5800. Burton, S. A., and J. I. Prosser. 2001. Autotrophic ammonia oxidation at low pH through urea hydrolysis. Appl. Environ. Microbiol. 67:2952–2957. Campbell, G. R. 2000. Community Analysis of Betasubgroup Ammonia-oxidising Bacteria in Sewage Sludge Amended Soil (PhD thesis). University of Aberdeen. Aberdeen, UK. Casciotti, K. L., and B. B. Ward. 2001. Dissimilatory nitrite reductase genes from autotrophic ammonia-oxidizing bacteria. Appl. Environ. Microbiol. 67:2213–2221. Ceccherini, M. T., M. Castialdini, C. Piovanelli, R. C. Hastings, A. J. McCarthy, M. Bazzicalupoet, and N. Miclaus. 1998. Effects of swine manure on autotrophic ammonia-

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

oxidizing bacteria in soil microcosms. Appl. Soil Ecol. 7:149–157. Chandler, D. P., R. W. Schreckhise, J. L. Smith, and H. Bolton. 1997. Electroelution to remove humic compounds from soil DNA and RNA extracts. J. Microbiol. Meth. 28:11– 19. Chang, Y. J., A. K. Hussain, J. R. Stephen, M. D. Mullen, D. C. White, and A. Peacock. 2001. Impact of herbizides on the abundance and structure of indigenous betasubgroup ammonia-oxidizer communities in soil microcosms. Environ. Toxicol. Chem. 20:2462–2468. Daims, H., J. L. Nielsen, P. H. Nielsen, K. H. Schleifer, and M. Wagner. 2001a. In situ characterization of Nitrospira-like nitrite-oxidizing bacteria active in wastewater treatment plants. Appl. Environ. Microbiol. 67:5273– 5284. Daims, H., U. Purkhold, L. Bjerrum, E. Arnold, P. A. Wilderer, and M. Wagner. 2001b. Nitrification in sequencing biofilm batch reactors: Lessons from molecular approaches. Water Sci. Tech. 43:9–18. Daims, H., N. B. Ramsing, K. H. Schleifer, and M. Wagner. 2001c. Cultivation-independent, semiautomatic determination of absolute bacterial cell numbers in environmental samples by fluorescence in situ hybridization. Appl. Environ. Microbiol. 67:5810–5818. de Bie, M. J. M., A. G. C. L. Speksnijder, G. A. Kowalchuk, T. Schuurmann, G. Zwart, J. R. Stephen, O. E. Diekmann, and H. J. Laanbroek. 2001. Shifts in the dominant populations of ammonia-oxidizing β-subclass Proeteobacteria along the eutrophic Schelde estuary. Aquat. Microb. Ecol. 23:225–236. De Boer, W., and H. J. Laanbroek. 1989. Ureolytic nitrification at low pH by Nitrosospira species. Arch. Microbiol. 152:178–181. De Boer, W., P. J. A. Klein-Gunnewiek, M. Veenhuis, E. Bock, and H. J. Laanbroek. 1991. Nitrification at low pH by aggregated autotrophic bacteria. Appl. Environ. Microbiol. 57:3600–3604. Dionisi, H. M., A. C. Layton, G. Harms, I. R. Gregory, K. G. Robinson, and G. S. Sayler. 2002. Quantification of Nitrosomonas oligotropha-like ammonia-oxidizing bacteria and Nitrospira spp. from full-scale wastewater treatment plants by competitive PCR. Appl. Environ. Microbiol. 68:245–253. Feray, C., B. Volat, V. V. Degrange, A. Clays-Josserand, and B. Montuelle. 1999. Assessment of three methods for detection and quantification of nitrite-oxidizing bacteria and nitrobacter in freshwater sediments (MPN-PCR, MPN-Griess, immunofluorescence). Microb. Ecol. 37:208–217. Gieseke, A., U. Purkhold, M. Wagner, R. Amann, and A. Schramm. 2001. Community structure and activity dynamics of nitrifying bacteria in a phosphate-removing biofilm. Appl. Environ. Microbiol. 67:1351–1362. Golovacheva, R. S. 1976. Thermophilic nitrifying bacteria from hot springs. Microbiology 45:329–331. Goreau, T. W., W. A. Kaplan, S. C. Wofsy, M. B. McElroy, F. W. Valois, and S. W. Watson. 1980. Production of NO2− and N2O by nitrifying bacteria at reduced concentrations of oxygen. Appl. Environ. Microbiol. 40:526–532. Guschin, D. Y., B. K. Mobarry, D. Proudnikov, D. A. Stahl, B. E. Rittmann, and A. D. Mirzabekov. 1997. Oligonucleotide microchips as genosensors for determinative and environmental studies in microbiology. Appl. Environ. Microbiol. 63:2397–2402.

807

Hall, G. H. 1986. Nitrification in lakes. In: J. I. Prosser (Ed.) Nitrification. IRL press. Washington, DC. 127–156. Harms, H., H.-P. Koops, and H. Wehrmann. 1976. An ammonia-oxidizing bacterium, Nitrosovibrio tenuis nov. gen. nov. sp. Arch. Microbiol. 108:105–111. Hastings, R. C., M. T. Ceccherini, N. Miclaus, J. R. Saunders, M. Bazzicalupo, and A. J. McCarthy. 1997. Direct molecular biological analysis of ammonia oxidising bacteria populations in cultivated soil plots treated with swine manure. FEMS Microbiol. Ecol. 23:45–54. Head, I. M., W. D. Hiorns, T. M. Embley, A. J. McCarthy, and J. R. Saunders. 1993. The phylogeny of autotrophic ammonia-oxidizing bacteria as determined by analysis of 16S ribosomal RNA gene sequences. J. Gen. Microbiol. 139:1147–1153. Helmer, C., S. Kunst, S. Juretschko, M. C. Schmid, K.-H. Schleifer, and M. Wagner. 1999. Nitrogen loss in a nitrifying biofilm system. Water Sci. Tech. 39:13–21. Henckel, T., M. Friedrich, and R. Conrad. 1999. Molecular analyses of the methane-oxidizing microbial community in rice field soil by targeting the genes of the 16S rRNA, particulate methane monooxygenase, and methanol dehydrogenase. Appl. Environ. Microbiol. 65:1980– 1990. Hermansson, A., and P. E. Lindgren. 2001. Quantification of ammonia-oxidizing bacteria in arable soil by real-time PCR. Appl. Environ. Microbiol. 67:972–976. Hiorns, W. D., R. C. Hastings, I. M. Head, A. J. McCarthy, J. R. Saunders, R. W. Pickup, and G. H. Hall. 1995. Amplification of 16S ribosomal RNA genes of autotrophic ammonia-oxidizing bacteria demonstrates the ubiquity of nitrosospiras in the environment. Microbiology 141:2793–800. Hollibaugh, J. T., N. Bano, and H. W. Ducklow. 2002. Widespread distribution in polar oceans of a 16S rRNA gene sequence with affinity to Nitrosospira-like ammoniaoxidizing bacteria. Appl. Environ. Microbiol. 68:1478– 1484. Holmes, A. J., A. Costello, M. E. Lidstrom, and J. C. Murrell. 1995. Evidence that particulate methane monooxygenase and ammonia monooxygenase may be evolutionarily related. FEMS Microbiol. Lett. 132:203–208. Holmes, A. J., P. Roslev, I. R. McDonald, N. Iversen, K. Henriksen, and J. C. Murrell. 1999. Characterization of methanotrophic bacterial populations in soils showing atmospheric methane uptake. Appl. Environ. Microbiol. 65:3312–3318. Hommes, N. G., L. A. Sayavedra-Soto, and D. J. Arp. 1998. Mutagenesis and expression of amo, which codes for ammonia monooxygenase in Nitrosomonas europaea. J. Bacteriol. 180:3353–3359. Horz, H. P., J. H. Rotthauwe, T. Lukow, and W. Liesack. 2000. Identification of major subgroups of ammonia-oxidizing bacteria in environmental samples by T-RFLP analysis of amoA PCR products. J. Microbiol. Meth. 39:197–204. Hoshino, T., N. Noda, S. Tsuneda, A. Hirata, and Y. Inamori. 2001. Direct detection by in situ PCR of the amoA gene in biofilm resulting from a nitrogen removal process. Appl. Environ. Microbiol. 67:5261–5266. Hovanec, T. A., and E. F. DeLong. 1996. Comparative analysis of nitrifying bacteria associated with freshwater and marine aquaria. Appl. Environ. Microbiol. 62:2888– 2896. Ivanova, I. A., J. R. Stephen, Y. J. Chang, J. Bruggemann, P. E. Long, J. P. McKinley, G. A. Kowalchuk, D. C. White, and S. J. Macnaughton. 2000. A survey of 16S rRNA and

808

H.-P. Koops et al.

amoA genes related to autotrophic ammonia-oxidizing bacteria of the beta-subdivision of the class Proteobacteria in contaminated groundwater. Can. J. Microbiol. 46:1012–1020. Jiang, Q. Q., and L. R. Bakken. 1999. Comparison of Nitrosospira strains isolated from terrestrial environments. FEMS Microbiol. Ecol. 30:171–186. Johnstone, B. H., and R. D. Jones. 1988. Physiological effects of long-term energy-source deprivation on the survival of a marine chemolithotrophic ammonia-oxidizing bacterium. Mar. Ecol. Prog. Ser. 49:295–303. Jones, R. D., R. Y. Morita, H.-P. Koops, and S. W. Watson. 1988. A new marine ammonium-oxidizing bacterium, Nitrosomonas cryotolerans sp. nov. Can. J. Microbiol. 34:1122–1128. Juretschko, S., G. Timmermann, M. Schmid, K. H. Schleifer, A. Pommerening-Roser, H. P. Koops, and M. Wagner. 1998. Combined molecular and conventional analyses of nitrifying bacterium diversity in activated sludge: Nitrosococcus mobilis and Nitrospira-like bacteria as dominant populations. Appl. Environ. Microbiol. 64:3042–3051. Klotz, M. G., and J. M. Norton. 1995. Sequence of an ammonia monooxygenase subunit A-encoding gene from Nitrosospira sp. NpAV. Gene 163:159–160. Koops, H.-P., H. Harms, and H. Wehrmann. 1976. Isolation of a moderate halophilic ammonia–oxidizing bacterium, Nitrosococcus mobilis nov. sp. Arch. Microbiol. 10:277– 282. Koops, H.-P., and H. Harms. 1985. Deoxyribonucleic acid homologies among 96 strains of ammonia-oxidizing bacteria. Arch. Microbiol. 141:214–218. Koops, H.-P., B. Böttcher, U. C. Möller, A. PommereningRöser, and G. Stehr. 1990. Description of a new species of Nitrosococcus. Arch. Microbiol. 154:244–248. Koops, H.-P., B. Böttcher, U. C. Möller, A. PommeringRöser, and G. Stehr. 1991. Classification of eight new species of ammonia-oxidizing bacteria: Nitrosomonas communis sp. nov., Nitrosomonas ureae sp. nov., Nitrosomonas aestuarii sp. nov., Nitrosomonas marina sp. nov., Nitrosomonas nitrosa sp. nov., Nitrosomonas eutropha sp. nov., Nitrosomonas oligotropha sp. nov., and Nitrosomonas halophila sp. nov. J. Gen. Microbiol. 137:1689–1699. Koops, H.-P., and U. C. Möller. 1992. The lithotrophic ammonia-oxidizing bacteria. In: A. Balows, H. G. Trü per, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York, NY. 2625–2637. Koops, H.-P., and A. Pommerening-Röser. 2001. Distribution and ecophysiology of the nitrifying bacteria emphasizing cultured species. FEMS Microbiol. Ecol. 37:1–9. Kowalchuk, G. A., J. R. Stephen, W. De Boer, J. I. Prosser, T. M. Embley, and J. W. Woldendorp. 1997. Analysis of ammonia-oxidizing bacteria of the beta subdivision of the class Proteobacteria in coastal sand dunes by denaturing gradient gel electrophoresis and sequencing of PCR-amplified 16S ribosomal DNA fragments. Appl. Environ. Microbiol. 63:1489–1497. Kowalchuk, G. A., Z. S. Naoumenko, P. J. L. Derikx, A. Felske, J. R. Stephen, and H. J. Laanbroek. 1998. Community analysis of ammonia-oxidising bacteria, in relation to oxygen availability in soils and root-oxygenated sediments, using PCR, DGGE and oligonucleotide probe hybridisation. FEMS Microbiol. Ecol. 27:339– 350.

CHAPTER 3.2.12 Kowalchuk, G. A., A. W. Stienstra, G. H. Heilig, J. R. Stephen, and J. W. Woldendorp. 2000a. Changes in the community structure of ammonia-oxidizing bacteria during secondary succession of calcareous grasslands. Environ. Microbiol. 2:99–110. Kowalchuk, G. A., A. W. Stienstra, G. H. Heilig, J. R. Stephen, and J. W. Woldendorp. 2000b. Molecular analysis of ammonia-oxidising bacteria in soil of successional grasslands of the Drentsche A (The Netherlands). FEMS Microbiol. Ecol. 31:207–215. Kowalchuk, G. A., and J. R. Stephen. 2001. Ammoniaoxidizing bacteria: A model for molecular microbial ecology. Ann. Rev. Microbiol. 55:485–529. Lee, N., P. H. Nielsen, K. H. Andreasen, S. Juretschko, J. L. Nielsen, K.-H. Schleifer, and M. Wagner. 1999. Combination of fluorescent in situ hybridization and microautoradiography: A new tool for structure-function analyses in microbial ecology. Appl. Environ. Microbiol. 65:1289–1297. Liebig, T., M. Wagner, L. Bjerrum, and M. Denecke. 2001. Nitrification performance and nitrifier community composition of a chemostat and a membrane-assisted bioreactor for the nitrification of sludge reject water. Bioproc. Biosyst. Engin. 24:203–210. Logemann, S., J. Schantl, S. Bijvank, M. van Loosdrecht, J. G. Kuenen, and M. Jetten. 1998. Molecular microbial diversity in a nitrifying reactor system without sludge retention. FEMS Microbiol. Ecol. 27:239–249. Ludwig, W., O. Strunk, S. Klugbauer, N. Klugbauer, M. Weizenegger, J. Neumaier, M. Bachleitner, and K.-H. Schleifer. 1998. Bacterial phylogeny based on comparative sequence analysis. Electrophoresis 19:554–568. MacDonald, R. M. 1986. Nitrification in soil: An introductory history. In: J. I. Prosser (Ed.) Nitrification. IRL Press. Oxford, UK. 1–16. McCaig, A. E., T. M. Embley, and J. I. Prosser. 1994. Molecular analysis of enrichment cultures of marine ammonia oxidisers. FEMS Microbiol. Lett. 120:363–367. McCaig, A. E., C. J. Phillips, J. R. Stephen, G. A. Kowalchuk, S. M. Harvey, R. A. Herbert, T. M. Embley, and J. I. Prosser. 1999. Nitrogen cycling and community structure of proteobacterial beta-subgroup ammonia-oxidizing bacteria within polluted marine fish farm sediments. Appl. Environ. Microbiol. 65:213–220. McTavish, H., J. A. Fuchs, and A. B. Hooper. 1993. Sequence of the gene coding for ammonia monooxygenase in Nitrosomonas europaea. J. Bacteriol. 175:2436–2444. Mendum, T. A., R. E. Sockett, and P. R. Hirsch. 1999. Use of molecular and isotopic techniques to monitor the response of autotrophic ammonia-oxidizing populations of the beta subdivision of the class Proteobacteria in arable soils to nitrogen fertilizer. Appl Environ. Microbiol. 65:4155–4162. Migula, W. 1900. System der Bakterien 2:194–195. Mobarry, B. K., M. Wagner, V. Urbain, B. E. Rittmann, and D. A. Stahl. 1996. Phylogenetic probes for analyzing abundance and spatial organization of nitrifying bacteria [published erratum appears in Appl. Environ. Microbiol.(1997), 63(2), p. 815]. Appl. Environ. Microbiol. 62:2156–2162. Morgenroth, E., A. Obermayer, E. Arnold, A. Brü hl, M. Wagner, and P. A. Wilderer. 2000. Effect of long-term idle periods on the performance of sequencing batch reactors. Water Sci. Tech. 41:105–113. Morris, S. A., S. Radajewski, T. W. Willison, and J. C. Murrell. 2002. Identification of the functionally active methan-

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

otroph population in a peat soil microcosm by stableisotope probing. Appl. Environ. Microbiol. 68:1446– 1453. Murray, R. G. E., and S. W. Watson. 1965. Structure of Nitrosocystis oceanus and comparison with Nitrosomonas and Nitrobacter. J. Bacteriol. 89:1594–1609. Nicolaisen, M. H., and N. B. Ramsing. 2002. Denaturing gradient gel electrophoresis (DGGE) approaches to study the diversity of ammonia-oxidizing bacteria. J. Microbiol. Meth. 50:189–203. Nogueira, R., L. F. Melo, U. Purkhold, S. Wuertz, and M. Wagner. 2002. Nitrifying and heterotrophic population dynamics in biofilm reactors: Effects of hydraulic retention time and the presence of organic carbon. Water Res. 36:469–481. Nold, S. C., J. Zhou, A. H. Devol, and J. M. Tiedje. 2000. Pacific Northwest marine sediments contain ammonia-oxidizing bacteria in the beta subdivision of the Proteobacteria. Appl. Environ. Microbiol. 66:4532– 4535. Norton, J. M., J. J. Alzerreca, Y. Suwa, and M. G. Klotz. 2002. Diversity of ammonia monooxygenase operon in autotrophic ammonia-oxidizing bacteria. Arch. Microbiol. 177:139–149. Okabe, S., H. Satoh, and Y. Watanabe. 1999. In situ analysis of nitrifying biofilms as determined by in situ hybridization and the use of microelectrodes. Appl. Environ. Microbiol. 65:3182–3191. Oved, T., A. Shaviv, T. Goldrath, R. T. Mandelbaum, and D. Minz. 2001. Influence of effluent irrigation on community composition and function of ammonia-oxidizing bacteria in soil. Appl. Environ. Microbiol. 67:3426– 3433. Painter, H. A. 1986. Nitrification in the treatment of sewage and waste waters. In: J. I. Prosser (Ed.) Nitrification. IRL Press. Oxford, UK. 185–211. Pedersen, K., J. Arlinger, L. Hallback, and C. Petersson. 1996. Investigations of subterranean bacteria in deep crystalline bedrock and their importance for the disposal of nuclear waste. Can. J. Microbiol. 42:382–391. Phillips, C. J., Z. Smith, T. M. Embley, and J. I. Prosser. 1999. Phylogenetic differences between particle-associated and planktonic ammonia-oxidizing bacteria of the beta subdivision of the class Proteobacteria in the Northwestern Mediterranean Sea. Appl. Environ. Microbiol. 65:779–786. Phillips, C. J., D. Harris, S. L. Dollhopf, K. L. Gross, J. I. Prosser, and E. A. Paul. 2000. Effects of agronomic treatments on structure and function of ammoniaoxidizing communities. Appl. Environ. Microbiol. 66:5410–5408. Pinck, C., C. Coeur, P. Potier, and E. Bock. 2001. Polyclonal antibodies recognizing the AmoB protein of ammonia oxidizers of the beta-subclass of the class Proteobacteria. Appl. Environ. Microbiol. 67:118–124. Pommerening-Röser, A. 1993. Untersuchungen zur Phylogenie Ammoniak oxidierender Bakterien (PhD thesis). University of Hamburg. Hamburg, Germany. Pommerening-Röser, A., G. Rath, and H.-P. Koops. 1996. Phylogenetic diversity within the genus Nitrosomonas. Syst. Appl. Microbiol. 19:344–351. Princic, A., I. I. Mahne, F. Megusar, E. A. Paul, and J. M. Tiedje. 1998. Effects of pH and oxygen and ammonium concentrations on the community structure of nitrifying bacteria from wastewater. Appl. Environ. Microbiol. 64:3584–3590.

809

Purkhold, U., A. Pommerening-Röser, S. Juretschko, M. C. Schmid, H.-P. Koops, and M. Wagner. 2000. Phylogeny of all recognized species of ammonia oxidizers based on comparative 16S rRNA and amoA sequence analysis: Implications for molecular diversity surveys. Appl. Environ. Microbiol. 66:5368–5382. Purkhold, U., M. Wagner, G. Timmermann, A. PommereningRöser, and H.-P. Koops. 2003. 16S rRNA and amoAbased phylogeny of 12 novel betaproteobacterial ammonia oxidizing isolates: Extension of the data set and proposal of a new lineage within the Nitrosomonads. Int. J. Syst. Evol. Microbiol. 53:1485–1494. Radajewski, S., G. Webster, D. S. Reay, S. A. Morris, P. Ineson, D. B. Nedwell, J. I. Prosser, and J. C. Murrell. 2002. Identification of active methylotroph populations in an acidic forest soil by stable-isotope probing. Microbiology 148:2331–2342. Radeva, G., K. Flemming, and S. Selenska–Pobell. 1999. Molecular analysis of bacterial populations in ground water pollutes with heavy metals. In: G. Bernhard (Ed.) Annual Report, Institute of Biochemistry. Institute of Biochemistry. 57. Rath, G. 1996. PhD thesis. University of Hamburg. Hamburg, Germany. Regan, J. M., G. W. Harrington, and D. R. Noguera. 2002. Ammonia- and nitrite-oxidizing bacterial communities in a pilot-scale chloraminated drinking water distribution system. Appl. Environ. Microbiol. 68:73–81. Rotthauwe, J. H., W. de Boer, and W. Liesack. 1995. Comparative analysis of gene sequences encoding ammonia monooxygenase of Nitrosospira sp. AHB1 and Nitrosolobus multiformis C-71. FEMS Microbiol. Lett. 133:131– 135. Rotthauwe, J. H., K. P. Witzel, and W. Liesack. 1997. The ammonia monooxygenase structural gene amoA as a functional marker: molecular fine-scale analysis of natural ammonia-oxidizing populations. Appl. Environ. Microbiol. 63:4704–4712. Sakano, Y., K. D. Pickering, P. F. Strom, and L. J. Kerkhof. 2002. Spatial distribution of total, ammonia-oxidizing, and denitrifying bacteria in biological wastewater treatment reactors for bioregenerative life support. Appl. Environ. Microbiol. 68:2285–2293. Schmid, M., U. Twachtmann, M. Klein, M. Strous, S. Juretschko, M. Jetten, J. W. Metzger, K.-H. Schleifer, and M. Wagner. 2000. Molecular evidence for a genuslevel diversity of bacteria capable of catalyzing anaerobic ammonium oxidation. Syst. Appl. Microbiol. 23:93– 106. Schmidt, I., and E. Bock. 1997. Anaerobic ammonia oxidation with nitrogen dioxide by Nitrosomonas eutropha. Arch. Microbiol. 167:106–111. Schmidt, I., and E. Bock. 1998. Anaerobic ammonia oxidation with nitrogen dioxide by cell-free extracts of Nitrosomonas eutropha. Ant. v. Leeuwenhoek 73:271–278. Schramm, A., L. H. Larsen, N. P. Revsbech, N. B. Ramsing, R. Amann, and K.-H. Schleifer. 1996. Structure and function of a nitrifying biofilm as determined by in situ hybridization and the use of microelectrodes. Appl. Environ. Microbiol. 62:4641–4647. Schramm, A., D. De Beer, M. Wagner, and R. Amann. 1998. Identification and activities in situ of Nitrosospira and Nitrospira spp. as dominant populations in a nitrifying fluidized bed reactor. Appl. Environ. Microbiol. 64:3480–5.

810

H.-P. Koops et al.

Schramm, A., D. de Beer, J. C. van den Heuvel, S. Ottengraf, and R. Amann. 1999. Microscale distribution of populations and activities of Nitrosospira and Nitrospira spp. along a macroscale gradient in a nitrifying bioreactor: quantification by in situ hybridization and the use of microsensors. Appl. Environ. Microbiol. 65:3690–3696. Schramm, A., D. De Beer, A. Gieseke, and R. Amann. 2000. Microenvironments and distribution of nitrifying bacteria in a membrane-bound biofilm. Environ. Microbiol. 2:680–6. Silyn-Roberts, G., and G. Lewis. 2001. In situ analysis of Nitrosomonas spp. in wastewater treatment wetland biofilms. Water Res. 35:2731–2739. Sinigalliano, C. D., D. N. Kuhn, and R. D. Jones. 1995. Amplification of the amoA gene from diverse species of ammonium-oxidizing bacteria and from an indigenous bacterial population from seawater. Appl. Environ. Microbiol. 61:2702–2706. Smith, Z., A. E. McCaig, J. R. Stephen, T. M. Embley, and J. I. Prosser. 2001. Species diversity of uncultured and cultured populations of soil and marine ammonia oxidizing bacteria. Microb. Ecol. 42:228–237. Smorzewski, W. T., and E. I. Schmidt. 1991. Numbers, activities, and diversity of autotrophic ammonia-oxidizing bacteria in a freshwater, eutrophic lake sediment. Can. J. Microbiol. 37:828–833. Sorokin, D. Y., G. Muyzer, T. Brinkhoff, J. G. Kuenen, and M. S. M. Jetten. 1998. Isolation and characterization of a novel facultatively alkaliphilic Nitrobacter species, N. alkalicus sp. nov. Arch. Microbiol. 170:345– 352. Sorokin, D., T. Tourova, M. C. Schmid, M. Wagner, H. P. Koops, J. G. Kuenen, and M. Jetten. 2001. Isolation and properties of obligately chemolithoautotrophic and extremely alkali-tolerant ammonia-oxidizing bacteria from Mongolian soda lakes. Arch. Microbiol. 176:170– 177. Speksnijder, A. G., G. A. Kowalchuk, K. Roest, and H. J. Laanbroek. 1998. Recovery of a Nitrosomonas-like 16S rDNA sequence group from freshwater habitats. Syst. Appl. Microbiol. 21:321–330. Speksnijder, A. G., G. A. Kowalchuk, S. De Jong, E. Kline, J. R. Stephen, and H. J. Laanbroek. 2001. Microvariation artifacts introduced by PCR and cloning of closely related 16S rRNA gene sequence. Appl. Environ. Microbiol. 67:469–472. Spieck, E., M. Meincke, and E. Bock. 1992. Taxonomic diversity of Nitrosovibrio strains isolated from building stones. FEMS Microbiol. Ecol. 102:21–26. Stackebrandt, E., R. G. E. Murray, and H. G. Trü per. 1988. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “purple bacteria and their relatives”. Int. J. Syst. Bacteriol. 38:321–325. Stackebrandt, E., and B. M. Goebel. 1994. Taxonomic note: a place for DNA-DNA reassociation and 16S rRNA sequence analysis in the present species definition in bacteriology. Int. J. Syst. Bacteriol. 44:846–849. Stehr, G., B. Böttcher, P. Dittberner, G. Rath, and H. P. Koops. 1995a. The ammonia-oxidizing nitrifying population of the River Elbe estuary. FEMS Microbiol. Ecol. 17:177–186. Stehr, G., S. Zörner, B. Böttcher, and H.-P. Koops. 1995b. Exopolymers: An ecological characteristic of a flocattached, ammonia-oxidizing bacterium. Microb. Ecol. 30:115–126.

CHAPTER 3.2.12 Stephen, J. R., A. E. McCaig, Z. Smith, J. I. Prosser, and T. M. Embley. 1996. Molecular diversity of soil and marine 16S rRNA gene sequences related to β-subgroup ammonia-oxidizing bacteria. Appl. Environ. Microbiol. 62:4147–4154. Stephen, J. R., G. A. Kowalchuk, M. A. V. Bruns, A. E. McCaig, C. J. Phillips, T. M. Embley, and J. I. Prosser. 1998. Analysis of beta-subgroup proteobacterial ammonia oxidizer populations in soil by denaturing gradient gel electrophoresis analysis and hierarchical phylogenetic probing. Appl. Environ. Microbiol. 64:2958–2965. Stephen, J. R., Y. J. Chang, S. J. Macnaughton, G. A. Kowalchuk, K. T. Leung, C. A. Flemming, and D. C. White. 1999. Effect of toxic metals on indigenous soil betasubgroup proteobacterium ammonia oxidizer community structure and protection against toxicity by inoculated metal-resistant bacteria. Appl. Environ. Microbiol. 65:95–101. Suwa, Y., Y. Imamura, T. Suzuki, T. Tashiro, and Y. Urushigawa. 1994. Ammonia-oxidizing bacteria with different sensitivities to (NH4 )2 SO4 in activated sludge. Water Res. 28:1523–1532. Suwa, Y., T. Sumino, and K. Noto. 1997. Phylogenetic relationships of activated sludge isolates of ammonia oxidizers with different sensitivities to ammonium sulfate. J. Gen. Appl. Microbiol. 43:373–379. Szwerinski, H., S. Gaiser, and D. Bardtke. 1985. Immunofluorescence for the quantitative determination of nitrifying bacteria: Interference of the test in biofilm reactors. Appl. Microbiol. Biotechnol. 21:125–128. Takahashi, R., N. Kondo, K. Usui, T. Kanehira, M. Shinohara, and T. Tokuyama. 1992. Pure isolation of a new chemoautotrophic ammonia-oxidizing bacterium on a gellan gum plate. J. Ferment. Bioeng. 74:52–54. Teske, A., E. Alm, J. M. Regan, T. S., B. E. Rittmann, and D. A. Stahl. 1994. Evolutionary relationships among ammonia- and nitrite-oxidizing bacteria. J. Bacteriol. 176:6623–6630. Tokuyama, T., N. Yoshida, T. Matasuishi, N. Takahashi, T. Takahashi, T. Kanehira, and M. Shinohara. 1997. A new psychrotrophic ammonia oxidizing bacterium Nitrosovibrio sp. TYM9. J. Ferment. Bioeng. 83:377–380. Trü per, H. G., and L. de Clari. 1997. Taxonomic note: Necessary correction of specific epithets formed as substantives (nouns) “in apposition”. Int. J. Syst. Bacteriol. 47:908–909. Utaaker, J. B., L. Bakken, Q. Q. Jiang, and I. F. Nes. 1995. Phylogenetic analysis of seven new isolates of ammoniaoxidizing bacteria based on 16S rRNA gene sequences. Syst. Appl. Microbiol. 18:549–559. Utaaker, J. B., and I. F. Nes. 1998. A qualitative evaluation of the published oligonucleotides specific for the 16S rRNA gene sequences of the ammonia-oxidizing bacteria. Syst. Appl. Microbiol. 21:72–88. Völsch, A., W. F. Nader, H. K. Geiss, G. Nebe, and C. Birr. 1990. Detection and analysis of two serotypes of ammonia-oxidizing bacteria in sewage plants by flow cytometry. Appl. Environ. Microbiol. 56:2430–2435. Voytek, M. A., and B. B. Ward. 1995. Detection of ammonium-oxidizing bacteria of the beta-subclass of the class Proteobacteria in aquatic samples with the PCR. Appl. Environ. Microbiol. 61:1444–1450. Voytek, M. A. 1996. Relative Abundance and Species Diversity of Autotrophic Ammonia Oxidizing Bacteria in Aquatic Systems (PhD thesis). University of California at Santa Cruz. Reston, Virginia.

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

Wagner, M., G. Rath, R. Amann, H.-P. Koops, and K.-H. Schleifer. 1995. In situ identification of ammoniaoxidizing bacteria. Syst. Appl. Microbiol. 18:251–264. Ward, B. B., and M. J. Perry. 1980. Immunofluoreszent assay for the marine ammonium-oxidizing bacterium Nitrosomonas oceanus. Appl. Environ. Microbiol 39:913–918. Ward, B. B. 1982. Oceanic distribution of ammoniumoxidizing bacteria determined by immunofluorscent assay. J. Mar. Res. 40:1155–1172. Ward, B. B., and A. F. Carlucci. 1985. Marine ammonia- and nitrit-oxidizing bacteria: serological diversity determined by immunofluorescence in sewage plants by flow cytometry. Appl. Environ. Microbiol. 50:194–201. Ward, B. B., M. A. Voytek, and K. Witzel. 1997. Phylogenetic diversity of natural populations of ammonia oxidizers investigated by specific PCR amplification. Microb. Ecol. 33:87–96. Ward, B. B., D. P. Martino, M. C. Diaz, and S. B. Joye. 2000. Analysis of ammonia-oxidizing bacteria from hypersaline Mono Lake, California, on the basis of 16S rRNA sequences. Appl. Environ. Microbiol. 66:2873– 2881. Ward, B. B., and G. D. O’Mullan. 2002. Worldwide distribution of Nitrosococcus oceani, a marine ammoniaoxidizing gamma-proteobacterium, detected by PCR and sequencing of 16S rRNA and amoA genes. Appl. Environ. Microbiol. 68:4153–4157. Watson, S. W. 1965. Characteristics of a marine nitrifying bacterium, Nitrosocystis oceanus sp. n. Limnol. Oceanogr. 10 (Suppl.):R274–R289. Watson, S. W., and C. C. Remsen. 1970. Cell envelope of Nitrosocystis oceanus. J. Ultrastr. Res. 33:148–160. Watson, S. W. 1971a. Taxonomic considerations of the family Nitrobacteraceae Buchanan: Requests for opinions. Int. J. Syst. Bacteriol. 21:254–270. Watson, S. W., and M. Mandel. 1971b. Comparison of the morphology and deoxyribonucleid acid composition of 27 strains of nitrifying bacteria. J. Bacteriol. 107:563– 569. Watson, S. W., L. B. Graham, C. C. Remsen, and F. W. Valois. 1971c. A lobular, ammonia oxidizing bacterium, Nitrosolobus multiformis, nov. sp. Arch. Microbiol. 76:183–203. Watson, S. W., F. W. Valois, and J. B. Waterbury. 1981. The family Nitrobacteraceae. In: M. P. Starr, H. Stolp, and H. Trü per (Eds.) The Prokaryotes. Springer-Verlag. Berlin, Germany. 1:1005–1022. Watson, S. W., E. Bock, H. Harms, H.-P. Koops, and A. B. Hooper. 1989. Nitrifying bacteria. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 3:1808–1834. Webster, G., T. M. Embley, and J. I. Prosser. 2002. Grassland management regimens reduce small-scale heterogeneity and species diversity of beta-proteobacterial ammonia oxidizer populations. Appl. Environ. Microbiol. 68:20– 30.

811

Whitby, C. B., J. R. Saunders, J. Rodriguez, R. W. Pickup, and A. McCarthy. 1999. Phylogenetic differentiation of two closely related Nitrosomonas spp. that inhabit different sediment environments in an oligotrophic freshwater lake. Appl. Environ. Microbiol. 65:4855– 4862. Whitby, C. B., G. Hall, R. Pickup, J. R. Saunders, P. Ineson, N. R. Parekh, and A. McCarthy. 2001a. 13C incorporation into DNA as a means of identifying the active components of ammonia-oxidizer populations. Lett. Appl. Microbiol. 32:398–401. Whitby, C. B., J. R. Saunders, R. W. Pickup, and A. J. McCarthy. 2001b. A comparison of ammonia-oxidiser populations in eutrophic and oligotrophic basins of a large freshwater lake. Ant. v. Leeuwenhoek 79:179–188. Wilhelm, R., A. Abeliovich, and A. Nejidat. 1998. Effect of long-term ammonia starvation on the oxidation of ammonia and hydroxylamine by Nitrosomonas europaea. J. Biochem. 124:811–815. Winogradsky, S. 1890. Recherches sur les organismes de la nitrification. Ann. Inst. Pasteur 4:213–331. Winogradsky, S. 1892. Contributions a la morphologie des organismes de la nitrification. Arch. Sci. Biol. (St. Petersburg) 1:88–137. Winogradsky, S., and H. Winogradsky. 1933. Etudes sur la microbiologie du sol. VII: Nouvelles recherches sur les organismes de la nitrification. Ann. Inst. Pasteur 50. Winogradsky, H. 1937. Contribution a l’étude de la microflore nitrificatrice des boues activées de Paris. Ann. Inst. Pasteur 58:326–340. Wintzigerode, F., and U. B. Goebel. 1997. Determination of microbial diversity in environmental samples: Pitfalls of PCR-based analysis. FEMS Microbiol. Rev. 21:213– 229. Woese, C. R., W. G. Weisburg, B. J. Paster, C. M. Hahn, R. S. Tanner, N. R. Krieg, H.-P. Koops, H. Harms, and E. Stackebrandt. 1984. The phylogeny of the purple bacteria: the beta subdivision. Syst. Appl. Microbiol. 5:327– 336. Woese, C. R., W. G. Weisburg, C. M. Hahn, B. J. Paster, L. B. Zablen, B. J. Lewis, T. J. Macke, W. Ludwig, and E. Stackebrandt. 1985. The phylogeny of the purple bacteria: The gamma subdivision. Syst. Appl. Microbiol. 6:25– 33. Wu, L., D. K. Thompson, G. Li, R. A. Hurt, J. M. Tiedje, and J. Zhou. 2001. Development and evaluation of functional gene arrays for detection of selected genes in the environment. Appl. Environ. Microbiol. 67:5780– 5790. Yamagata, A., J. Kato, R. Hirota, A. Kuroda, T. Ikeda, N. Takiguchi, and H. Ohtake. 1999. Isolation and characterization of two cryptic plasmids in the ammoniaoxidizing bacterium Nitrosomonas sp. strain ENI-11. J. Bacteriol. 181:3375–3381. Zart, D., I. Schmidt, and E. Bock. 2000. Significance of gaseous NO for ammonia oxidation by Nitrosomonas eutropha. Ant. v. Leeuwenhoek 77:49–55.

Prokaryotes (2006) 5:812–827 DOI: 10.1007/0-387-30745-1_37

CHAPTER 3.2.13 ehT

suneG

su l l i cabo i hT

The Genus Thiobacillus LESLEY A. ROBERTSON AND J. GIJS KUENEN

Introduction

Habitats

Ever since the genus Thiobacillus was first described in 1904, the ability to grow while using a reduced sulfur compound as a source of energy has been considered sufficiently important taxonomically to merit classifying all Gram-negative, sulfur-oxidizing, nonphototrophic rods in this genus. However, as studies using modern taxonomic methods began to reveal that some of the species are only superficially related, virtually every paper describing a new Thiobacillus species in the last decade has mentioned the need to reorganize the genus. Most of this reorganization has now been done (Kelly and Wood, 2000a), and the authors are grateful to D.P. Kelly and A. Wood for permission to see and use their resulting manuscripts prior to publication. The effect of the reorganization is dramatic (Table 1). In the last edition of The Prokaryotes, the genus Thiobacillus included 17 species. There are now only three species left—all autotrophs, and all members of the β-subclass of the Proteobacteria. The 4 or 5 facultative autotrophs that fall into the β-subclass have now been assigned to a new genus—Thiomonas. Most of the other former Thiobacillus species fall into the γ-subclass, together with the members of the genus Thiomicrospira. This includes both the well-known acidiphilic and recently discovered alkaliphilic “Thiobacillus” species, as well as the marine species. However, because the reorganization is still so new and most researchers will look for these organisms in one group, many of them will be dealt with in this chapter, with the exception of the γ group. Thiobacillus acidophilus and Thiobacillus versutus have both been transferred to existing genera (see Table 1). Similarly, Thiosphaera pantotropha has now been reclassified as Paracoccus pantotrophus and will be described in the chapter on that genus. The last member of the γ group, Thiobacillus novellus, is now Starkeya novella. Within this chapter, the generic terms “thiobacilli” and “colorless sulfur bacteria” refer to the whole group, regardless of their new classification.

In nature, the distribution of reduced inorganic sulfur compounds is only one of the factors governing the presence and activity of the colorless sulfur bacteria, and a complex of chemical, physical, and microbial interactions is almost as important a controlling force. One crucial factor is the requirement for the simultaneous presence of an electron donor (the reduced sulfur compound) and electron acceptor (i.e., oxygen or nitrogen oxides). Sulfide is one of the most important natural substrates, and many of the colorless sulfur bacteria grow in the narrow zones and gradients where sulfide and oxygen coexist (Nelson and Jannasch, 1983). Such gradients can be found, for example, in stratified lakes and at the interface between aerobic water and an anaerobic sediment, but may be even more common in microniches in environments that contain anaerobic pockets (e.g., sediments and wet soils). It must be realized that extremely high turnover rates of sulfide can be observed at very low levels of sulfide and oxygen (3,000 Da) hemolytic activity was described in one strain of B. cepacia that was able to induce nucleosomal degradation (consistent with apoptosis) in human neutrophils and in a mouse macrophage cell line (Hutchison et al., 1998). There does not appear to be any correlation between production of extracellular virulence factors and transmissibility of epidemic B. cepacia isolates. Some strains of the B. cepacia complex have been shown to invade and survive in cultured epithelial cells and macrophages (Burns et al., 1996; Martin and Mohr, 2000; Saini et al., 1999). Burkholderia cepacia also has been reported to be resistant to growth inhibition by epithelial cell β-defensins and resistant to nonoxidative neutrophil-killing mechanisms compared to P. aeruginosa (Baird et al., 1999; Speert et al., 1994). The ability to acquire iron from host sources is a prerequisite for the successful establishment and maintenance of most bacterial infections. Yang et al. have demonstrated that 84 : 84 B. pseudomallei strains examined during their studies tested positive for siderophore production using the chrome azurol S (CAS) assay. A structural and chemical analysis of the siderophore synthesized by B. pseudomallei U7 confirmed the molecule was approximately 1,000 Da in size, was water soluble with a yellow-green fluorescence, and belonged to the hydroxamate class (Yang et al., 1991). Furthermore, studies also have demonstrated the siderophore was capable of scavenging iron from both lactoferrin and

CHAPTER 3.2.16

transferrin in vitro (Yang et al., 1993). The name malleobactin has been proposed for this compound (Yang et al., 1991). Although B. pseudomallei isolates can express an impressive array of both secreted and cellassociated antigens, the role(s) of these products in the pathogenesis of disease to date have been relatively ill defined. One of the primary reasons for this has been the lack of suitable techniques for genetically manipulating the organism. Owing to the recent application of a transposon-based mutagenesis system for use in B. pseudomalle, genetic loci, which encode a number of these putative virulence determinants and protective antigens, have been identified and characterized (DeShazer et al., 1997; DeShazer et al., 1998). It has been shown previously that B. pseudomallei isolates can secrete antigens that demonstrate biological activities consistent with proteases, lecithinases, lipases and hemolysins (Esselman and Liu, 1961; Ashdown and Koehler, 1990; Sexton et al., 1994). However, though the importance of these factors has been implicated in the pathogenesis of the disease, only the protease has been characterized to date. Studies conducted by Sexton et al. (1994) have confirmed the presence of a 36-kDa antigen with associated proteolytic activities in B. pseudomallei culture supernatants. In particular, a protease expressed by B. pseudomallei (isolate 319a) was found to be a metalloenzyme requiring iron for maximal protease activity and to be optimally active at pH 8.0 and 60°C (Sexton et al., 1994). Furthermore, monoclonal antibodies (MAb) raised against a Pseudomonas aeruginosa alkaline protease were cross-reactive with this antigen (Sexton et al., 1994). Most recently, an 11.8-kb chromosomal locus in B. pseudomallei has been identified that demonstrates a high degree of homology to operons that encode for the products of the main terminal branch of the general secretory pathway (GSP; Pugsley, 1993). Further characterization of the open-reading frames in this locus have confirmed that their orientation and physical arrangement are virtually identical to the pul gene cluster of Klebsiella oxytoca (Pugsley, 1993). Not surprisingly, the phenotypic analysis of individual transposon mutants also has confirmed their inability to secrete antigens associated with protease, lipase and lecithinase into the extracellular milieu. Interestingly, although the protease, lipase and lecithinase may play a small role in the pathogenesis of acute melioidosis, mutants deficient in their ability to secrete these particular exoenzymes were not severely attenuated in their ability to cause a fulminating illness (DeShazer et al., 1999). In the mid-1950s, several studies demonstrated that filter-sterilized B. pseudomallei culture

CHAPTER 3.2.16

supernatants were lethal for mice and hamsters when administered parenterally (Nigg et al., 1955; Heckly and Nigg, 1958; Heckly, 1964). These results were consistent with the fulminating illnesses observed in animals following inoculation with viable bacteria and suggested that B. pseudomallei strains might be capable of secreting a lethal toxin. In studies conducted by Ismail et al. (1987), mouse lethal, thermolabile toxin was presumably purified to homogeneity and characterized as a 31-kDa protein. Haase et al. (1997) have also described the presence of cytotoxic activity in culture filtrates. Their results, however, suggest that the antigen is only 3 kDa in size and that the cytotoxic activity in this instance most likely is due to the presence of a small peptide. Recently it has been reported that a rhamnolipid purified from B. pseudomallei culture supernatants demonstrates a cytotoxic effect against HL60 and HeLa cell lines (Haubler et al., 1998). Because this activity can be neutralized by albumin, however, it is unlikely to be of consequence in the pathogenesis of B. pseudomallei infections. These results have not been reproduced in animal models, even when using preparations concentrated by lyophilization (Brett et al., 1997; Brett et al., 1998). Burkholderia pseudomallei strains can synthesize capsular antigens (Smith et al., 1987; Leelarasamee and Bovornkitti, 1989), and they may play an important role in the pathogenesis of melioidosis. Whereas in vitro studies have determined that encapsulated B. pseudomallei strains are as susceptible to phagocytic uptake by polymorphonuclear leukocytes (PMNs) as non-encapsulated variants, evidence tends to suggest that the presence of exopolysaccharide confers upon them the ability to resist the bactericidal effects of the phagolysosomal environment (Smith et al., 1987; Pruksachartvuthi et al., 1990). This feature of B. pseudomallei strains may help to explain why these organisms are able to remain latent in a host for as long as 26 years. Recently, Steinmetz et al. isolated and purified a high-molecular-weight capsular antigen (150 kDa) from B. pseudomallei NCTC 7431 and succeeded in raising a Mab against it (Steinmetz et al., 1995). They were able to demonstrate the reactivity of both mucoid and non-mucoid strains with the MAb, suggesting that the capsular antigen is constitutively expressed by B. pseudomallei strains. Interestingly, temperature appeared to have little effect on the synthesis of the exopolysaccharide inasmuch as B. pseudomallei strains grown at both 15 and 37°C were Mab-reactive. Furthermore, an assay utilizing a variety of Pseudomonas spp. and Burkholderia spp. as controls was able to

The Genus Burkholderia

855

confirm the specificity of the MAb for B. pseudomallei and B. mallei strains only (Steinmetz et al., 1995). More recently, Masoud et al. (1997) have been successful at elucidating the chemical and structural characteristics of a capsular polysaccharide isolated from the virulent clinical isolate B. pseudomallei 304b. Their results demonstrated that the exopolysaccharide was a linear unbranched polymer of repeating tetrasaccharide units having the structure (-3)-2-O-Ac-β-D-Galp-(1-4)-α-D-Galp-(1-3)-β-DGalp-(1-5)-β-D-KDOp-(2-) (where KDO is 3deoxy-D-manno-2-octulosonic acid). Similarly, Nimtz et al. (1997) have demonstrated that a structurally identical capsular antigen is expressed by B. pseudomallei NCTC 7431. Studies by both groups have also shown that patient sera reacted strongly with the purified carbohydrate antigens, indicating that this carbohydrate polymer is most likely expressed in vivo (Steinmetz et al., 1995; Masoud et al., 1997). Previous studies have confirmed that the lipopolysaccharide (LPS) antigens expressed by B. pseudomallei strains are highly conserved throughout this species (Pitt et al., 1992). In fact, serological evidence suggests only one serotype of B. pseudomallei (Bryan et al., 1994). To investigate this phenomenon, Perry et al. have characterized the LPS antigens isolated from a number of B. pseudomallei strains (Perry et al., 1995). Their results demonstrated that B. pseudomallei strains coordinately express two distinct somatic O-antigens (PS) on their cell surface. The type I antigen consists of a highmolecular-weight unbranched 1,3-linked homopolymer of 2-O-acetylated 6-deoxy-β-Dmanno-heptopyranosyl residues, while the type II antigen is an unbranched heteropolymer consisting of (-3)-β-D-glucopyranose-(1-3)-6deoxy-α-L-talopyranosyl-(1)-disaccharide repeats (L-6dTalp: __33% O-4 acetylated and O-2 methylated; __66% O-2 acetylated; Knirel et al., 1992; Perry et al., 1995). Even though the simultaneous expression of two or more LPS moieties is not an uncommon feature associated with Gram-negative bacteria, the degree to which the two PS antigens are conserved amongst B. pseudomallei strains is quite remarkable (Perry et al., 1995). It has been reported that B. pseudomallei strains are resistant to the bactericidal effects of normal human serum (Ismail et al., 1988). Type II PS is essential for conferring this resistance phenotype. A 17.5-kb region of the chromosome, which is required for the synthesis of the type II antigen, confers the serum-resistance phenotype (DeShazer et al., 1998). Type II PS is probably a significant determinant in the pathogenesis of melioidosis because the LD50 value associated with a type II PS mutant is approximately 140-

856

D.E. Woods and P.A. Sokol

fold higher than that of the wild-type strain (Woods et al., 1993; DeShazer et al., 1998). Flagella are commonly recognized as important virulence determinants expressed by bacterial pathogens because the motility phenotype imparted by these organelles often correlates with the ability of an organism to cause disease (Penn and Luke, 1992; Moens and Vanderleyden, 1996). A significant degree of size and antigenic homogeneity exists amongst flagellins expressed by B. pseudomallei isolates. Furthermore, flagellin-specific antiserum can passively protect diabetic infant rats against a B. pseudomallei challenge (Brett et al., 1994). Curiously, there was no significant difference between the virulence capacities associated with a wild-type strain of B. pseudomallei and nonmotile mutants in either the diabetic infant rat or Syrian hamster models of infection (DeShazer et al., 1997). These results indicate that though flagella and/or motility may not be major virulence determinants in the pathogenesis of melioidosis, purified flagellin might still be a protective immunogen against B. pseudomallei infections. Although glanders is a serious life-threatening zoonotic disease, relatively little is known about the pathogenesis, virulence factors, strain and the host immunopathological responses to infection (Fritz et al., 1999).

Treatment Burkholderia cepacia is resistant to most antibiotics, making treatment of CF patients very difficult. It usually is susceptible only to piperacillin, axlocillin, cefoperazone, ceftazidime, chloramphenicol and trimethoprim-sulfamethoxazole (TMP-SMX; Pitkin et al., 1997). Strains isolated from CF sputum from patients treated with several courses of antibiotic therapy are often resistant to all known antimicrobial agents (Gilligan and Whittier, 1999). Resistance to a variety of antimicrobial agents including penicillins, firstand second-generation cephalosporins and many of the aminoglycosides is characteristic of B. pseudomallei clinical isolates (Dance et al., 1988; Leelarasamee and Bovornkitti, 1989; Godfrey et al., 1991; Weinberg and Heller, 1997). With this in mind, accurate identification of the organism, evaluation of the severity of the infection and antibiotic susceptibility testing are of paramount importance in devising an effective chemotherapeutic strategy. Whereas the newer therapies that utilize combinations of ceftazidime–cotrimoxazole or amoxicillin–clavulanate for treatment of disease are proving beneficial, the mortality rates associated with the acute septicemic and pulmonary forms of melioidosis are still unacceptably high (Smith et al., 1987;

CHAPTER 3.2.16

Leelarasamee and Bovornkitti, 1989; Kanai and Kondo, 1994; Weinberg and Heller, 1997; Ho et al., 1997). Typically, prolonged oral therapy also is recommended to assure the full clinical resolution of infections while reducing the potential for recrudescence of disease. Antibiotic therapy for glanders is complex (Batmanov, 1991); however, there have been reports that ceftazidime, imipenem, doxycycline and ciprofloxacin are active against both B. mallei and B. pseudomallei (Kenny et al., 1999). Antibiotics clinically proven to be effective in the treatment of melioidosis may therefore be effective for treating glanders. Acknowledgements. This work was supported by Department of Defense Contract DAMD 17-98-C-8003, the Canadian Bacterial Diseases Network of Centres of Excellence, the Medical Research Council of Canada and the Canadian Cystic Fibrosis Foundation.

Literature Cited Ashdown, L. R., and J. M. Koehler. 1990. Production of hemolysin and other extracellular enzymes by clinical isolates of Pseudomonas pseudomallei. J. Clin. Microbiol. 28:2331–2334. Baird, R. M., H. Brown, A. W. Smith, and M. L. Watson. 1999. Burkholderia cepacia is resistant to the antimicrobial activity of airway epithelial cells. Immunopharmacology 44(3):267–272. Batmanov, V. P. 1991. Sensitivity of Pseudomonas mallei to fluoroquinolones and their efficacy in experimental glanders. Antibiot. Khimioter. 36:31–34. Bauernfeind, A., C. Roller, D. Meyer, R. Jungwirth, and I. Schneider. 1998. Molecular procedure for rapid detection of Burkholderia mallei and Burkholderia pseudomallei. J Clin Microbiol. 36(9):2737–2744. Bauernfeind, A., I. Schneider, R. Jungwirth, and C. Roller. 1999. Discrimination of Burkholderia multivorans and Burkholderia vietnamiensis from Burkholderia cepacia genomovars I, III and IV by PCR. J. Clin. Microbiol. 37:1335–1339. Benenson, A. S. 1995. Control of Communicable Diseases manual. American Public Health Association. Washington DC, 577. Bharadwaj, R., A. Kagal, S. K. Deshpandy, S. A. Joshi, P. M. Khare, A. R. Junnakar, and M. A. Phadke. 1994. Outbreak of plague-like illness caused by Pseudomonas pseudomallei in Maharashtra, India. Lancet 344:1574. Brett, P. J., D. C. Mah, and D. E. Woods. 1994. Isolation and characterization of Pseudomonas pseudomallei flagellin proteins. Infect. Immunol. 62:1914–1919. Brett, P. J., D. DeShazer, and D. E. Woods. 1997. Characterization of Burkholderia pseudomallei and Burkholderia pseudomallei-like strains. Epidemiol. Infect. 118:137– 148. Brett, P. J., D. DeShazer, and D. E. Woods. 1998. Burkholderia thailandensis sp. Nov., a Burkhoderia pseudomalleilike species. Int. J. System. Bacteriol. 48:317–320.

CHAPTER 3.2.16 Bryan, L. E., S. Wong, D. E. Woods, D. A. B. Dance, and W. Chaowagul. 1994. Passive protection of diabetic rats with antisera specific for the polysaccharide portion of the lipopolysaccharide isolated from Pseudomonas pseudomallei. Can. J. Infect. Dis. 5:170–178. Burkholder, W. H. 1950. Sour skin, a bacterial rot of onion bulbs. Phytopathology 40:115–117. Burns, J. L., M. Jonas, E. Y. Chi, D. K. Clark, A. Berger, and A. Griffith. 1996. Invasion of respiratory epithelial cells by Burkholderia (Pseudomonas) cepacia. Infect. Immunol. 64(10):4054–4059. Chaowagul, W., N. J. White, D. A. B. Dance, Y. Wattanagoon, P. Naigowit, T. M. E. Davis, S. Looareesuwan, and N. Pitawatchara. 1989. Melioidosis, a major cause of community-acquired septicemia in northeastern Thailand. J. Infect. Dis. 159:890–899. Corey, M., and V. Farewell. 1996. Determinants of mortality from cystic fibrosis in Canada, 1970–1989. Am. J. Epidemiol. 143(10):1007–1017. Crosa, J. H. 1989. Genetics and molecular biology of siderophore-mediated iron transport in bacteria. Microbiol. Rev. 53(4):517–530. Dance, D. A., V. Wuthiekanun, N. J. White, and W. Chaowagul. 1988. Antibiotic resistance in Pseudomonas pseudomallei. Lancet 1:994–995. Dance, D. A., V. Wuthiekanun, P. Naigowit, and N. J. White. 1989. Identification of Pseudomonas pseudomallei in clinical practice: use of simple screening tests and API 20NE. J. Clin. Pathol. 42:645–648. Dance, D. A. 1990. Melioidosis. Rev. Med. Microbiol. 1:143– 150. Dance, D. A. 1991. Melioidosis: The tip of the iceberg? Clin. Microbiol. Rev. 4:52–60. Darling, P., M. Chan, A. D. Cox, and P. A. Sokol. 1998. Siderophore production by cystic fibrosis isolates of Burkholderia cepacia. Infect. Immunol. 66(2):874–877. DeShazer, D., P. Brett, R. Carlyon, and D. E. Woods. 1997. Mutagenesis of Burkholderia pseudomallei with Tn 5OT182: isolation of motility mutants and molecular characterization of the flagellin structural gene. J. Bacteriol. 179:2116–2125. DeShazer, D., P. Brett, and D. E. Woods. 1998. The Type II O-antigen moiety of Burkholderia pseudomallei lipopolysaccharide is required for serum resistance and virulence. Molec. Microbiol. 30:1011–1081. Deshazer, D., and D. E. Woods. 1999. Pathogenesis of melioidosis: Use of Tn5-OT182 to study the molecular basis of Burkholderia pseudomallei virulence. J. Infect. Dis. Antimicrob. Ag. 16:91–96. Ellison, D. W., H. J. Baker, and M. Mariappan. 1969. Melioidosis in Malaysia. I: A method of isolation of Pseudomonas pseudomallei from soil and surface water. Am. J. Trop. Med. Hyg. 18:694–697. Esselman, M. T., and P. V. Liu. 1961. Lecithinase production by Gram-negative bacteria. J. Bacteriol. 81:939–945. Frangolias, D. D., E. Mahenthiralingam, S. Rae, J. M. Raboud, A. G. F. Davidson, R. Wittmann, and P. G. Wilcox. 1999. Burkholderia cepacia in cystic fibrosis: Variable disease course. Am. J. Respir. Crit. Care Med. 160:1572–1577. Freeman, B. A. 1985. Pseudomonas and Legionella. In: Burrow’s Textbook of Microbiology. W. B. Saunders Co. Philadelphia, PA. 544–557. Fritz, D. L, P. Vogel, D. R. Brown, and D. M. Waag. 1999. The hamster model of intraperitoneal Burkholderia mallei (glanders). Vet. Pathol. 36(4):276–291.

The Genus Burkholderia

857

Gilligan, P. H., P. A. Gage, L. M. Bradshaw, D. V. Schidlow, and B. T. DeCicco. 1985. Isolation medium for the recovery of Pseudomonas cepacia from respiratory secretions of patients with cystic fibrosis. J. Clin. Microbiol. 22:5–8. Gilligan, P. H. 1991. Microbiology of airway disease in patients with cystic fibrosis. Clin. Microbiol. Rev. 4(1):35–51. Gilligan, P. H., and S. Whittier. 1999. Burkholderia, Stenotrophomonas, Ralstonia, Brevundimonas, Comamonas, and Acidovorax. In: P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (Eds.) Manual of Clinical Microbiology. ASM Press. Washington DC. Gillis, M., T. V. Van, R. Bardin, M. Goor, P. Hebbar, A. Willems, P. Segers, K. Kersters, T. Heulin, and M. P. Fernandez. 1995. Polyphasic taxonomy in the genus Burkholderia leading to an amended description of the genus and proposition of Burkholderia vietnamiensis sp. nov. for N2-fixing isolates from rice in Vietnam. Int. J. Syst. Bacteriol. 45:274–289. Godfrey, A. J., S. Wong, D. A. Dance, W. Chaowagul, and L. E. Bryan. 1991. Pseudomonas pseudomallei resistance to beta-lactam antibiotics due to alterations in the chromosomally encoded beta-lactamase. Antimicrob. Agents Chemother. 35:1635–1640. Goldman, D. A., and J. D. Klinger. 1986. Pseudomonas cepacia: Biology, mechanisms of virulence, epidemiology. J. Pediatr. 108:806–812. Govan, J. R., P. H. Brown, J. Maddison, C. J. Doherty, J. W. Nelson, M. Dodd, A. P. Greening, and A. K. Webb. 1993. Evidence for transmission of Pseudomonas cepacia by social contact in cystic fibrosis. Lancet 342(8862):15–19. Govan, J. R., and V. Deretic. 1996a. Microbial pathogenesis in cystic fibrosis: Mucoid Pseudomonas aeruginosa and Burkholderia cepacia. Microbiol. Rev. 60(3):539–574. Govan, J. R., J. E. Hughes, and P. Vandamme. 1996b. Burkholderia cepacia: Medical, taxonomic and ecological issues. J. Med. Microbiol. 45(6):395–407. Haase, A., J. Janzen, S. Barrett, and B. Currie. 1997. Toxin production by Burkholderia pseudomallei strains and correlation with severity of melioidosis. J. Med. Microbiol. 46:557–563. Haubler, S., M. Nimtz, T. Domke, V. Wray, and I. Steinmetz. 1998. Purification and characterization of a cytotoxic exolipid of Burkholderia pseudomallei. Infect. Immunol. 66:1588–1593. Heckly, R. J., and C. Nigg. 1958. Toxins of Pseudomonas pseudomallei. II: Characterization. J. Bacteriol. 76:427– 436. Heckly, R. J. 1964. Differentiation of exotoxin and other biologically active substances in Pseudomonas pseudomallei filtrates. J. Bacteriol. 88:1730–1736. Henry, D. A., M. E. Campbell, J. J. LiPuma, and D. P. Speert. 1997. Identification of Burkholderia cepacia isolates from patients with cystic fibrosis and use of a simple new selective medium. J. Clin. Microbiol. 35:614–619. Henry, D. A., M. Campbell, C. McGimpsey, A. Clarke, L. Louden, J. Burns, M. H. Roe, P. Vandamme, and D. Speert. 1999. Comparison of isolation media for recovery of Burkholderia cepacia complex from respiratory secretions of patients with cystic fibrosis. J. Clin. Microbiol. 37:1004–1007. Ho, M., T. Schollaardt, M. D. Smith, M. B. Perry, P. J. Brett, W. Chaowagul, and L. E. Bryan. 1997. Specificity and functional activity of anti-Burkholderia pseudomallei polysaccharide antibodies. Infect. Immunol. 65:3648– 3653.

858

D.E. Woods and P.A. Sokol

Holmes, A., R. Nolan, R. Taylor, R. Finley, M. Riley, R. Z. Jiang, S. Steinbach, and R. Goldstein. 1999. An epidemic of Burkholderia cepacia transmitted between patients with and without cystic fibrosis. J. Infect. Dis. 179(5):1197–1205. Hornick, R. B. 1982. Diseases due to Pseudomonas mallei and Pseudomonas pseudomallei. In: R. J. Wedgewood (Ed.) Infections in Children. Harper & Row. Philadelphia, PA. 910–913. Howe, C. 1949. Glanders. In: H. A. Christian (Ed.) The Oxford Medicine. Oxford University Press. New York, NY. 185–201. Howe, C., A. Sampath, and M. Spotnitz. 1971. The Pseudomallei group: A review. J. Infect. Dis. 124:598– 606. Hutchison, M. L., I. R. Poxton, and J. R. Govan. 1998. Burkholderia cepacia produces a hemolysin that is capable of inducing apoptosis and degranulation of mammalian phagocytes. Infect. Immunol. 66(5):2033–2039. Inglis, T. J. J., S. C. Garrow, C. Adams, M. Henderson, and M. Mayo. 1998. Dry season outbreak of melioidosis in Western Australia. Lancet 352:1600. Isles, A., I. Maclusky, M. Corey, R. Gold, C. Prober, P. Fleming, and H. Levison. 1984. Pseudomonas cepacia infection in cystic fibrosis: An emerging problem. J. Pediatr. 104(2):206–210. Ismail, G., M. N. Embi, O. Omar, and N. Razak. 1987. Toxigenic properties of Pseudomonas pseudomallei extracellular products. Trop. Biomed. 4:101–110. Ismail, G., N. Razak, R. Mohamed, N. Embi, and O. Omar. 1988. Resistance of Pseudomonas pseudomallei to normal human serum bactericidal action. Microbiol. Immunol. 32:645–652. Johnson, W. M., S. D. Tyler, and K. R. Rozee. 1994. Linkage analysis of geographic and clinical clusters in Pseudomonas cepacia infections by multilocus enzyme electrophoresis and ribotyping. J. Clin. Microbiol. 32(4):924– 930. Kanai, K., and E. Kondo. 1994. Recent advances in biomedical sciences of Burkholderia pseudomallei (basonym: Pseudomonas pseudomallei). Jpn. J. Med. Sci. Biol. 47:1–45. Kenny, D. J., P. Russell, D. Rogers, S. M. Eley, and R. W. Titball. 1999. In vitro susceptibilities of Burkholderia mallei in comparison to those of other pathogenic Burkholderia spp.. Antimicrob. Agents Chemother. 43(11);Nov:2773–2775. Knirel, Y. A., N. A. Paramonov, A. S. Shashkov, N. K. Kochetkov, R. G. Yarullin, S. M. Farber, and V. I. Efremenko. 1992. Structure of the polysaccharide chains of Pseudomonas pseudomallei lipopolysaccharides. Carbohydr. Res. 233:185–193. Kooi, C., A. Cox, P. Darling, and P. A. Sokol. 1994. Neutralizing monoclonal antibodies to an extracellular Pseudomonas cepacia protease. Infect. Immunol. 62(7):2811–2817. Kovalev, G. K. 1971. Glanders (review). Zh. Mikrobiol. Edidemiol. Immunobiol. 48:63–70. Leelarasamee, A., and S. Bovornkitti. 1989. Melioidosis: Review and update. Rev. Infect. Dis. 11:413–425. Lewenza, S., B. Conway, E. P. Greenberg, and P. A. Sokol. 1999. Quorum sensing in Burkholderia cepacia: identification of the LuxRI homologs CepRI. J. Bacteriol. 181(3):748–756. LiPuma, J. J., J. E. Mortensen, S. E. Dasen, T. D. Edlind, D. V. Schidlow, J. L. Burns, and T. L. Stull. 1988.

CHAPTER 3.2.16 Ribotype analysis of Pseudomonas cepacia. J. Infect. Dis. 1164:133–136. LiPuma, J. J., S. E. Dasen, D. W. Neilson, R. C. Stern, and T. L. Stull. 1990. Person-to-person transmission of Pseudomonas cepacia between patients with cystic fibrosis. Lancet 336:527–532. LiPuma, J. J. 1998. Burkholderia cepacia: Management issues and new insights. Clin. Chest Med. 19(3):473–486, vi. Lonon, M. K., D. E. Woods, and D. C. Straus. 1988. Production of lipase by clinical isolates of Pseudomonas cepacia. J. Clin. Microbiol. 26(5):979–984. Mahenthiralingam, E., M. E. Campbell, D. A. Henry, and D. A. Speert. 1996. Epidemiology of Burkholderia cepacia infection in patients with cystic fibrosis: Analysis by random amplified polymorphic DNA fingerprinting. J. Clin. Microbiol. 34:2914–2920. Mahenthiralingam, E., D. A. Simpson, and D. P. Speert. 1997. Identification and characterization of a novel DNA marker associated with epidemic Burkholderia cepacia strains recovered from patients with cystic fibrosis. J. Clin. Microbiol. 35(4):808–816. Mahenthiralingam, E., T. Coenye, J. W. Chung, D. P. Speert, J. R. W. Govan, P. Taylor, and P. Vandamme. 2000. Diagnostically and experimentally useful panel of strains from the Burkholderia cepacia complex. J. Clin. Microbiol. 38(2):910–913. Martin, D. W., and C. D. Mohr. 2000. Invasion and intracellular survival of Burkholderia cepacia. Infect. Immunol. 68(1):24–29. Masoud, H., M. Ho, T. Schollaardt, and M. B. Perry. 1997. Characterization of the capsular polysaccharide of Burkholderia (Pseudomonas) pseudomallei 304b. J. Bacteriol. 179:5663–5669. Mays, E. E., and E. A. Ricketts. 1975. Melioidosis: Recrudescence associated with bronchogenic carcinoma twentysix years following initial geographic exposure. Chest 68:261–263. McCormick, J. B., D. J. Sexton, J. G. McMurray, E. Carey, P. Hayes, and R. A. Feldman. 1975. Human-to-human transmission of Pseudomonas pseudomallei. Ann. Internat. Med. 83:512–513. McKevitt, A. I., and D. E. Woods. 1984. Characterization of Pseudomonas cepacia isolates from patients with cystic fibrosis. J. Clin. Microbiol. 19(2):291–293. McKevitt, A. I., S. Bajaksouzian, J. D. Klinger, and D. E. Woods. 1989. Purification and characterization of an extracellular protease from Pseudomonas cepacia. Infect. Immunol. 57(3):771–778. Mietzner, T., and S. Morse. 1994. The role of iron-binding proteins in the survival of pathogenic bacteria. Ann. Rev. Nutr. 14:471–493. Moens, S. M., and J. Vanderleyden. 1996. Functions of bacterial flagella. Crit. Rev. Microbiol. 22:67–100. Mortensen, J. E., M. C. Fisher, and J. J. LiPuma. 1995. Recovery of Pseudomonas cepacia and other Pseudomonas species from the environment. Infect. Control. Hosp. Epidemiol. 16:30–32. Nakazawa, T., Y. Yamada, and M. Ishibashi. 1987. Characterization of hemolysin in extracellular products of Pseudomonas cepacia. J. Clin. Microbiol. 25(2):195– 198. Nigg, C., R. J. Heckly, and M. Colling. 1955. Toxin produced by Malleomyces pseudomallei. Proc. Soc. Exp. Biol. Med. 89:17–20. Nijhuis, E. H., M. J. Maat, I. W. E. Zeegers, C. Waalwijk, and J. A. Van-Venn, J.A. 1993. Selection of bacteria suitable

CHAPTER 3.2.16 for introduction into the rhizosphere of grass. Soil Biol. Biochem. 25:885–895. Nimtz, M., V. Wray, T. Domke, B. Brenneke, S. Haussler, and I. Steinmetz. 1997. Structure of an acidic exopolysaccharide of Burkholderia pseudomallei. Eur. J. Biochem. 250:608–616. Palleroni, N. J., and B. Holmes. 1981. Pseudomonas cepacia sp. nov., nom. rev. Int. J. Syst. Bacteriol. 31:479–481. Parker, M. T. 1990. Glanders and melioidosis. In: M. T. Parker and L. H. Collier (Eds.) Topley & Wilson’s Principles of Bacteriology, Virology and Immunity. B. C. Decker Inc. Philadelphia, PA. 392–394. Pegues, D. A., L. A. Carson, O. C. Tablan, S. C. FitzSimmons, S. B. Roman, J. M. Miller, and W. R. Jarvis. 1994. Acquisition of Pseudomonas cepacia at summer camps for patients with cystic fibrosis: Summer Camp Study Group. J. Pediatr. 124(5, Part 1):694–702. Pegues, C. F., D. F. Pegues, D. S. Ford, P. L. Hibberd, L. A. Carson, C. M. Raine, and D. C. Hooper. 1996. Burkholderia cepacia respiratory acquisition: epidemiology and molecular characterization of a large nosocomial outbreak. Epidemiol. Infect. 116:309–317. Penn, C. W., and C. J. Luke. 1992. Bacterial flagellar diversity and significance in pathogenesis. FEMS Microbiol. Lett. 100:331–336. Perry, M. B., L. L. MacLean, T. Schollaardt, L. E. Bryan, and M. Ho. 1995. Structural characterization of the lipopolysaccharide O antigens of Burkholderia pseudomallei. Infect. Immunol. 63:3348–3352. Pitkin, D. H., W. Sheikh, and H. L. Nadler. 1997. Comparative in vitro activity of meropenem versus other extended-spectrum antimicrobials against randomly chosen and selected resistant clinical isolates tested in 26 North American centers. Clin. Infect. Dis. 24(Suppl. 2):S238–S248. Pitt, T. L. 1990. Pseudomonas mallei and P. pseudomallei. In: M. T. Parker and L. H. Collier (Eds.) Topley & Wilson’s Principles of Bacteriology, Virology and Immunity. B. C. Decker Inc. Philadelphia, PA. 265–268. Pitt, T. L., H. Aucken, and D. A. Dance. 1992. Homogeneity of lipopolysaccharide antigens in Pseudomonas pseudomallei. J. Infect. 25:139–146. Prince, A. 1986. Antibiotic resistance of Pseudomonas species. J. Pediatr. 108:830–834. Pruksachartvuthi, S., N. Aswapokee, and K. Thakerngpol. 1990. Survival of Pseudomonas pseudomallei in human phagocytes. J. Med. Microbiol. 31:109–114. Pugsley, A. P. 1993. The complete general secretory pathway in Gram-negative bacteria. Microbiol. Rev. 57:50–108. Ralston, E., N. J. Palleroni, and M. Doudoroff. 1973. Pseudomonas pickettii, a new species of clinical origin related to Pseudomonas solanacearum. Int. J. Syst. Bacteriol. 23:15–19. Riboli, A. C., R. Koshinski, K. Arias, K. Marks-Austin, D. Stiertz, and T. L. Stull. 1996. An outbreak of Burkholderia cepacia lower respiratory tract infection associated with contaminated albuterol nebulization solution. Infect. Control Hosp. Epidemiol. 17:741–743. Saini, L. S., S. B. Galsworthy, M. A. John, and M. A. Valvano. 1999. Intracellular survival of Burkholderia cepacia complex isolates in the presence of macrophage cell activation. Microbiology 145(12):3465–3475. Sanford, J. P. 1990. Principles and Practice of Infectious Diseases. In: G. L. Mandell, R. G. Douglas Jr., and J. E. Bennett (Eds.) Churchill Livingstone. New York, NY. 1692–1696.

The Genus Burkholderia

859

Sanford, J. P. 1995. Pseudomonas species including melioidosis and glanders. In: G. L. Mandell, J. E. Bennett and R. Dolin (Eds.) Principles and Practice of Infectious Diseases, 4th ed. Churchill Livingstone. New York, NY. 1250–1254. Segonds, C., T. Heulin, N. Marty, and G. Chabanon. 1999. Differentiation of Burkholderia species by PCRrestriction fragment length polymorphism analysis of the 16S rRNA gene and application to cystic fibrosis isolates. J. Clin. Microbiol. 37:2201–2208. Sexton, M. M., A. L. Jones, W. Chaowagul, and D. E. Woods. 1994. Purification and characterization of a protease from Pseudomonas pseudomallei. Can. J. Microbiol. 40:903–910. Simpson, I. N., J. Finlay, D. J. Winstanley, N. Dewhurst, J. W. Nelson, S. L. Butler, and J. R. Govan. 1994. Multiresistance isolates possessing characteristics of both Burkholderia (Pseudomonas) cepacia and Burkholderia gladioli from patients with cystic fibrosis. J. Antimicrob. Chemother. 34(3):353–361. Smith, C. J., J. C. Allen, M. N. Embi, O. Othman, N. Razak, and G. Ismail. 1987. Human melioidosis: An emerging medical problem. MIRCEN J. 3:343–366. Smith, D. L., L. B. Gumery, E. G. Smith, D. E. Stableforth, M. E. Kaufmann, and T. L. Pitt. 1993. Epidemic of Pseudomonas cepacia in an adult cystic fibrosis unit: Evidence of person-to-person transmission. J. Clin. Microbiol. 31(11):3017–3022. Smith, M. D., V. Wuthiekanun, A. L. Walsh, and N. J. White. 1995. Quantitative recovery of Burkholderia pseudomallei from soil in Thailand. Trans. R. Soc. Trop. Med. Hyg. 89:488–490. Sokol, P. A. 1986. Production and utilization of pyochelin by clinical isolates of Pseudomonas cepacia. J. Clin. Microbiol. 23(3):560–562. Sokol, P. A., P. Darling, D. E. Woods, E. Mahenthiralingam, and C. Kooi. 1999. Role of ornibactin biosynthesis in the virulence of Burkholderia cepacia: characterization of pvdA, the gene encoding L-ornithine N(5)-oxygenase. Infect. Immunol. 67(9):4443–4455. Sokol, P. A., C. Kooi, R. S. Hodges, P. Cachia, and D. E. Woods. 2000. Immunization with a Pseudomonas elastase peptide reduces severity of experimental lung infections due to Pseudomonas aeruginosa or Burkholderia cepacia. J. Infect. Dis. 181:1682–1692. Speert, D. P., M. Bond, R. C. Woodman, and J. T. Curnutte. 1994. Infection with Pseudomonas cepacia in chronic granulomatous disease: Role of nonoxidative killing by neutrophils in host defense. J. Infect. Dis. 170(6):1524– 1531. Steele, J. H. 1973. The zoonoses: An epidemiologist’s viewpoint. In: M. Stefanini (Ed.) Progress in Clinical Pathology. Grune and Stratton. New York, NY. 239– 286. Steinmetz, I., M. Rohde, and B. Brenneke. 1995. Purification and characterization of an exopolysaccharide of Burkholderia (Pseudomonas) pseudomallei. Infect. Immunol. 63:3959–3965. Sun, L., R. Z. Jiang, S. Steinbach, A. Holmes, C. Campanelli, J. Forstner, U. Sajjan, Y. Tan, M. Riley, and R. Goldstein. 1995. The emergence of a highly transmissible lineage of cbl+ Pseudomonas (Burkholderia) cepacia causing CF centre epidemics in North America and Britain. Nat. Med. 1(7):661–666. Tanphaichitra, D., and S. Srimuang. 1984. Cellular immunity in tuberculosis, melioidoses, pasteurellosis, penicilliosis

860

D.E. Woods and P.A. Sokol

and role of levamisole and isoprinosine. Dev. Biol. Stand. 57:117–123. Tanphaichitra, D. 1989. Tropical disease in the immunocompromised host: melioidosis and pythiosis. Rev. Infect. Dis. 11(Suppl. 7):S1629–S1643. Vandamme, P., B. Holmes, M. Vancanneyt, T. Coenye, B. Hoste, R. Coopman, H. Revets, S. Lauwers, M. Gillis, K. Kersters, and J. R. Govan. 1997. Occurrence of multiple genomovars of Burkholderia cepacia in cystic fibrosis patients and proposal of Burkholderia multivorans sp. nov. Int. J. Syst. Bacteriol. 47(4):1188–1200. Vandamme, P., E. Mahenthiralingam, B. Holmes, T. Coenye, B. Hoste, P. De Vos, D. Henry, and D. P. Speert. 2000. Identification and population structure of Burkholderia stabilis sp. nov (formerly Burkholderia cepacia genomovar IV). J. Clin. Microbiol. 38:1042–1047. Von Graevenitz, A. 1973. Clinical microbiology of unusual Pseudomonas species. Prog. Clin. Path. 5: 185–218. Weinberg, A. N., and H. M. Heller. 1997. Infectious Diseases of the Lung. Thieme-Stratton. New York, NY. 2413– 2430. Welch, D. F., M. J. Muszynski, C. H. Pai, M. J. Maracon, M. M. Hribar, P. H. Gilligan, J. M. Matsen, P. A. Ahlin, B. C. Hilman, and S. A. Chartrand. 1987. Selective and differential medium for recovery of Pseudomonas cepacia from the respiratory tracts of patients with cystic fibrosis. J. Clin. Microbiol. 25:1730–1734. Whitmore, A., and C. S. Krishnaswami. 1912. An account of the discovery of a hitherto undescribed infective disease occurring among the population of Rangoon. Indian Med. Gaz. 47:262–267. Whitmore, A. 1913. An account of a glanders-like disease occurring in Rangoon. J. Hyg. Camb. 13:1–7.

CHAPTER 3.2.16 Woods, D. E., A. L. Jones, and P. J. Hill. 1993. Interaction of insulin with Pseudomonas pseudomallei. Infect. Immunol. 61:1914–1919. Wuthiekanum, V., M. D. Smith, D. A. B. Dance, A. L. Walsh, T. L. Pitt, and N. J. White. 1996a. Biochemical characteristics of clinical and environmental isolates of Burkholderia pseudomallei. J. Med. Microbiol. 45:408– 412. Wuthiekanum, V., M. D. Smith, D. A. B. Dance, A. L. Walsh, T. L. Pitt, and N. J. White. 1996b. Isolation of Pseudomonas pseudomallei from soil in north-eastern Thailand. Trans. R. Soc. Trop. Med. Hyg. 89:41–43. Yabuuchi, E., Y. Kosako, H. Oyaizu, I. Yano, H. Hotta, Y. Hashimoto, T. Ezaki, and M. Arakawa. 1992. Proposal of Burkholderia gen. nov. and transfer of seven species of the genus Pseudomonas homology group II to the new genus, with the type species Burkholderia cepacia (Palleroni and Holmes 1981) comb. nov. Microbiol. Immunol. 36(12):1251–1275. Yabuuchi, E., Y. Kosako, I. Yano, H. Hotta, and Y. Nishiuchi. 1995. Transfer of two Burkholderia and an Alcaligenes species to Ralstonia gen. nov.: Proposal of Ralstonia pickettii (Ralston, Palleroni and Doudoroff 1973) comb. nov., Ralstonia solanacearum (Smith 1896) comb. nov. and Ralstonia eutropha (Davis 1969) comb. nov. Microbiol. Immunol. 39:897–904. Yang, H. M., W. Chaowagul, and P. A. Sokol. 1991. Siderophore production by Pseudomonas pseudomallei. Infect. Immunol. 59:776–780. Yang, H. M., C. D. Kooi, and P. A. Sokol. 1993. Ability of Pseudomonas pseudomallei malleobactin to acquire transferrin-bound, lactoferrin-bound, and cell-derived iron. Infect. Immunol. 61:656–662.

Prokaryotes (2006) 5:861–872 DOI: 10.1007/0-387-30745-1_41

CHAPTER 3.2.17 ehT

e t i r t iN

gn i z i d i xO

a i re t caB

The Nitrite-Oxidizing Bacteria AHARON ABELIOVICH

Introduction Nitrite-oxidizing bacteria are a small group of primarily organo and/or chemoautotrophs that are difficult to grow and work with. For many years they drew very little attention, although it was realized that they provide a key link in the global nitrogen cycle between the ammoniaoxidizing bacteria, which generate nitrite, and the various denitrifying microorganisms that remove nitrate by reducing it to ammonia or molecular nitrogen, thus completing the global nitrogen cycle. The growing public awareness relating to issues of environmental pollution in recent decades brought renewed interest in this group of bacteria, as their role in the transformation of nitric oxides became apparent. In addition, these bacteria cause economic damage to agriculture and to ground water quality by contributing to the leaching of nitrogen fertilizers (distributed in the form of ammonia) from surface soils, thus polluting aquifers with increasing concentrations of nitrites and nitrates. Owing to their key role in the global cycling of nitrogen and their activities both as prime contributors to and scavengers of nitric oxides in the biosphere, nitrite-oxidizing bacteria became in recent years the focus of intensive ecophysiological research, in spite of the difficulties in cultivating and maintaining many of these bacteria in the laboratory. In addition to this group of autotrophic nitrifyers, there is also a diverse group of heterotrophic bacteria capable of heterotrophic nitrite oxidation. Thus, in one such study (Sakai et al., 1996), the nitrite transforming activities of heterotrophic bacterial strains from various culture collections as well as isolates from activated sludge were studied. Of the 48 strains tested, 17 strains consumed 1–5 mM of nitrite and accumulated corresponding amounts of nitrate. Heterotrophic microorganisms are rather flexible in determining the fate of the consumed nitrite. For instance, in Bacillus subtilis strain I-41 (a denitrification-positive isolate), the ratio of the amount

of nitrate accumulated to that of nitrite consumed varied from 0 to 100%, depending on the culture conditions. However, the mechanism of oxidation of nitrite to nitrate in these heterotrophic bacteria is very different from that in the autotrophic bacteria (Sorokin, 1991), as heterotrophic nitrite oxidation requires catalase and hydrogen peroxide (H2O2) generated through oxidation of organic electron donors. Also, aeration affects the nitrite- and nitrate-transforming activities of various heterotrophic bacteria. For example, Pseudomonas pavonaceae, a denitrification-positive strain, metabolizes both nitrite and nitrate to more reduced compounds at low oxygen pressure, and the direction of the conversion changes from reduction to oxidation at high oxygen pressure. This switching might be caused by inhibition and repression of the nitrite-reducing activity and by stimulation of nitrite-oxidizing activity by oxygen (Sakai et al., 1997).

Historic Background Nitrite-oxidizing bacteria carry the second stage of the nitrification process, that of oxidation of nitrite to nitrate. Because nitrate is an essential ingredient of gunpowder, nitrate and nitrification were the focus of interest both to scientists and politicians for many centuries. Nitrification protocols for the manufacture of nitrates by composting organic matter in soil have existed for many centuries: from the tenth century in China and from the twelfth century in Europe (Macdonald, 1986). However, the process was considered to be of a chemical and not biological nature until the late nineteenth century. It was only in 1862 that Pasteur suggested that nitrification was of biological origin (Pasteur, 1862), and it was only when Winogradsky in 1891 succeeded in isolating a nitrite-oxidizing bacterium that the debate about whether nitrification is a one-stage process (carried by a single bacterium) or a twostage process (carried by two distinct classes of microorganisms) was settled (Macdonald, 1986).

862

A. Abeliovich

CHAPTER 3.2.17

Nitrite Oxidation A detailed account of the mechanism of nitrite oxidation as an energy source has been described previously (Bock et al., 2001). The key enzyme that makes the nitrite-oxidizing bacteria so important in the global nitrogen cycle is nitrite oxidoreductase (NOR), which carries out the stoichiometric reaction: -

-

NO 2 + H 2O ¤ NO3 + 2H + + e -

(1)

2H + + 2e - + 0.5O 2 Æ H 2O

(2)

Reaction (1) is reversible, and many nitrite oxidizers are capable of reducing nitrate to nitrite under anaerobic conditions, but the significance of the reverse reaction to their survival is not clear, as both nitrite and nitrate are rapidly reduced in anaerobic environments. Nitrite oxidoreductase was isolated from various Nitrobacter strains, such as from mixotrophically grown cells of Nitrobacter hamburgensis; the enzyme purified from heat-treated membranes was homogeneous by the criteria of polyacrylamide gel electrophoresis and size exclusion chromatography. The monomeric form consisted of two subunits with molecular weights of 115 kDa and 65 kDa, respectively. The dimeric form of the enzyme contained 0.70 molybdenum, 23.0 iron, 1.76 zinc and 0.89 copper gram-atoms per molecule. The catalytically active enzyme was investigated by visible and electron paramagnetic resonance spectroscopy (EPR) under oxidizing (as isolated), reducing (dithionite), and turnover (nitrite) conditions. As isolated, the enzyme exhibited a complex set of EPR signals between 5–75 K, originating from several ironsulfur and molybdenum (V) centers. Addition of the substrate nitrite or the reducing agent dithionite resulted in a set of new resonances. The molybdenum and the iron-sulfur centers of nitrite oxidoreductase from Nitrobacter hamburgensis were involved in the transformation of nitrite to nitrate (Meincke et al., 1992).

Systematics As late as 1981, all bacteria capable of chemolithotrophic growth using ammonia ornitrite as energy source and capable of assimilating CO2 via the Calvin cycle were grouped in one family, the Nitrobacteraceae (Watson et al., 1981). This classification was based on the fact that all the bacteria grouped in this family carry out oxidation reactions of inorganic nitrogen compounds, and all are capable of chemolithotrophic growth using these as energy sources. All are Gramnegative bacteria and usually found in associ-

ation; as a physiological interdependence exists between the two groups, inasmuch as the ammonia oxidizers provide the substrate for the nitrite oxidizers, whereas the nitrite oxidizers are inhibited by excess ammonia. With the advent of molecular biology and classification of bacteria through sequencing of 16S ribosomal RNA genes and other specific DNA sequences, as well as the use of monoclonal antibodies, it became evident that the two groups are not related, and that there is no phylogenetic relationship between ammonia and nitrite oxidizers. Almost all nitrite oxidizers (with the exception of Nitrospira moscoviensis, which seems to occupy an intermediate position; Schramm et al., 1998) are a homogenous group belonging to the α-subdivision of the Proteobacteria and are very closely related to the nonsulfur purple photosynthetic bacterium Rhodopseudomonas palustris. None of the ammonia-oxidizing bacteria belong to the αsubdivision, and they are all distributed between the β- and γ-subdivisions (Woese et al., 1984; Woese et al., 1985). Comparative 16S rDNA sequencing was used to evaluate phylogenetic relationships among selected strains of ammonia- and nitriteoxidizing bacteria (Teske et al., 1994). All characterized strains were shown to be affiliated with the proteobacteria. The study extended recent 16S rDNA-based studies of phylogenetic diversity among nitrifiers by the comparison of eight strains of the genus Nitrobacter and representatives of the genera Nitrospira and Nitrospina. The latter genera were shown to be affiliated with the δ-subdivision of the proteobacteria but did not share a specific relationship to each other or to other members of the δ-subdivision. All characterized Nitrobacter strains constituted a closely related assemblage within the αsubdivision of the Proteobacteria. As previously observed, all ammonia-oxidizing genera except Nitrosococcus oceanus constitute a monophyletic assemblage within the β-subdivision of the proteobacteria. Consideration of physiology and phylogenetic distribution suggests that nitriteoxidizing bacteria of the α- and γ-subdivisions are derived from immediate photosynthetic ancestry. Each nitrifier retains the general structural features of the specific ancestor’s photosynthetic membrane complex. Thus, the nitrifiers, as a group, apparently are not derived from an ancestral nitrifying phenotype. Watson et al. (1981) identified three genera of nitrite-oxidizing bacteria (the Nitrospira being discovered only in 1986; Watson, 1986) and classified them on the basis of morphology and nutritional requirements. Bartosch et al. (1999) also divided the nitrite-oxidizing bacteria into four genera, Nitrobacter, Nitrospina, Nitrococcus and

CHAPTER 3.2.17

Nitrospira, according to their reactions with monoclonal antibodies prepared against the nitrite oxidoreductase of the genus Nitrobacter. Among these, Nitrobacter and Nitrospira strains are capable of heterotrophic or mixotrophic growth, the rest being strict chemolithotrophs. Because of the difficulties in culturing nitrifying bacteria on solid media, the methods for enumerating and identifying nitrifiers in environmental samples conventionally depended on incubating serial dilutions for very long periods, and then testing these for appearance of nitrate. Thus, the dominant nitrite-oxidizing species in an environmental sample was usually identified only after incubation of 3–6 months and was acertained on the basis of morphological and physiological properties such as sensitivity to salts and nutrient concentration. These observations led over the years to a general consensus that Nitrobacter strain were the dominant nitriteoxidizing organisms in practically all tested environments such as soil, marine waters, wastewater treatment plants, fresh water and reservoirs; Nitrococcus and Nitrospina were rarely identified (Watson et al., 1981). In 1995, a Gram-negative, nonmotile, nonmarine, nitrite-oxidizing bacterium was isolated by enrichment culture of a sample from a partially corroded area of an iron pipe located in a heating system in Moscow, Russia (Ehrich et al., 1995). The cells (0.9–2.2 µm × 0.2–0.4 µm in size) were helical to vibroid-shaped and often formed spirals with up to three turns 0.8–1.0 µm in width. The organism possessed an enlarged periplasmic space and lacked intracytoplasmic membranes and carboxysomes. The cells excreted extracellular polymers and formed aggregates. The bacterium grew optimally at 39°C and pH 7.6–8.0 in a mineral medium with nitrite as sole energy source and carbon dioxide as sole carbon source. The doubling time was 12 h in a mineral medium with 7.5 mM nitrite. The cell yield was low; only 0.9 mg of protein/liter was formed during oxidation of 7.5 mM nitrite. Under anoxic conditions, hydrogen could be used as electron donor with nitrate as electron acceptor. Organic matter (yeast extract, meat extract and peptone) supported neither mixotrophic nor heterotrophic growth. At concentrations as low as 0.75 g of organic matter/liter or higher, growth of the nitrite-oxidizing cells was inhibited. The cells contained cytochromes of the b- and c-type. The G+C content of DNA was 56.9 ± 0.4 mol%. This chemolithoautotrophic nitrite-oxidizer differed from the terrestrial members of the genus Nitrobacter with regard to morphology and substrate range and was similar to Nitrospira marina in both characteristics. The isolated bacterium was designated as a new species of the genus Nitrospira. Recent studies suggest that this genus

The Nitrite-Oxidizing Bacteria

863

dominates most of the nitrite oxidation activities in natural as well as artificial habitats (see below). With the introduction of molecular technologies to environmental microbiology, the true dominant nitrite oxidizers could be identified in situ in environmental samples, either by polymerase chain reaction (PCR, using specific rDNA primers) or fluorescence in situ hybridization (FISH, using specific probes). These studies led to the surprising conclusion that Nitrospira strains are the dominant nitrite oxidizers in most environmental samples tested so far, whereas Nitrobacter sp. are seldom identified without prior enrichment (Burrell et al., 1998; Burrell et al., 1999; Schramm et al., 1998; Juretschko et al.,1998; Okabe et al., 1999). There is, however, no doubt that within the next years, with the advent of oligonucleotide microchip technology, oligonucleotide microchips will dominate as genosensors for determinative and environmental studies in microbiology (Guschin et al., 1997). Precise identification of bacterial communities and of the genes switched on by each member of these communities under various environmental conditions will become routine in the environmental microbiology lab, and these procedures will enable the fine resolution of the community of nitrifying bacteria and its activities in different biotopes. DNA sequencing methodologies also were used to identify bacteria in situ, in biofilms and activated sludge particles (Schramm et al., 1998). Bacterial aggregates from a chemolithoautotrophic, nitrifying fluidized bed reactor were investigated with microsensors and rDNA-based molecular techniques. The microprofiles of O2, NH4+, NO2− and NO3− demonstrated the occurrence of complete nitrification in the outer 125 µm of the aggregates. FISH analysis showed that the dominant populations were of Nitrospira spp. and Nitrosospira sp. and that they formed separate, dense clusters which were in contact with each other and occurred throughout the aggregate. Significantly, no ammonia- or nitriteoxidizing bacteria of the genera Nitrosomonas or Nitrobacter, respectively, could be detected by FISH. To identify the nitrite oxidizers, a 16S ribosomal DNA clone library was constructed and screened by denaturing gradient gel electrophoresis (DGGE), and selected clones were sequenced. The organisms represented by these sequences formed two phylogenetically distinct clusters affiliated with the nitrite oxidizer Nitrospira moscoviensis. The dominant Nitrospira sp. formed clusters with the nitrosospira sp. and occurred throughout the aggregate while the second, smaller, morphologically and genetically different population of Nitrospira sp. was

864

A. Abeliovich

restricted to the outer nitrifying zones. The phylogeny of bacteria belonging to the genus Nitrobacter was investigated by sequencing the whole 16S rRNA gene (Orso et al., 1994). The average level of similarity for three Nitrobacter strains examined was high (99.2%), and the similarity level between Nitrobacter winogradsky and Nitrobacter sp. strain LL, which represent two different genomic species, was even higher (99.6%). When all of the Nitrobacter strains and their phylogenetic neighbors Bradyrhizobium and Rhodopseudomonas species were considered, the average similarity level was 98.1%. When complete sequences were used, Nitrobacter hamburgensis clustered with the two other Nitrobacter strains, though this was not the case when partial sequences were used. The two Rhodopseudomonas palustris strains examined exhibited a low similarity level (97.6%) and were not clustered. In another study (Okabe et al., 1999), the in situ spatial organization of ammonia-oxidizing and nitrite-oxidizing bacteria in domestic wastewater biofilms and autotrophic nitrifying biofilms was investigated by using microsensors and FISH, performed with 16S rDNA-targeted oligonucleotide probes. The combination of these techniques made it possible to relate in situ microbial activity directly to the occurrence of specific nitrifying bacterial populations. In situ hybridization revealed that bacteria belonging to the genus Nitrobacter were not detected; instead, Nitrospira-like bacteria were the main nitriteoxidizing bacteria in both types of biofilms. Nitrospira-like cells formed irregularly shaped aggregates consisting of small microcolonies, which bound the clusters of ammonia oxidizers. Whereas most of the ammonia-oxidizing bacteria were present throughout the biofilms, the nitrite-oxidizing bacteria were restricted to the active nitrite-oxidizing zones, which were inside the biofilms. Microelectrode measurements showed that the active ammonia-oxidizing zone was located in the outer part of a biofilm, whereas the active nitrite-oxidizing zone was located just below the ammonia-oxidizing zone and overlapped the location of nitrite-oxidizing bacteria, as determined by FISH. The use of fluorescent monoclonal antibodies was also very useful for the rapid quantification and in situ detection of specific nitrifiers in a mixed bacterial habitat such as a biofilm (Noda et al., 2000). In that study, 12 monoclonal antibodies against Nitrosomonas europaea (IFO14298) and 16 against Nitrobacter winogradsky (IFO 14297) enabled a direct cell count of N. europaea and N. winogradsky. Moreover, the distribution of N. europaea and N. winogradsky in a biofilm could be examined. Most of N. winogradsky existed near the surface and most

CHAPTER 3.2.17

of N. europaea existed within the polyethylene glycol (PEG) gel pellet. Using DNA sequences from the intergenic spacer (IGS) region of the ribosomal operon, and two primers derived from 16S and 23S rDNA conserved sequences, Navarro et al. (1992a) amplified these sequences by PCR. The PCR products, cleaved by four base cutting restriction enzymes, were used to differentiate Nitrobacter strains. This method is convenient for the genotypic characterization of Nitrobacter isolates and was successfully used to characterize natural populations of Nitrobacter from various soils and a lake. Considerable diversity was demonstrated in various soils, and in both water and sediments of the lake. Navarro et al. (1992b) also studied the genomic diversity of Nitrobacter strains by determining rDNA gene restriction patterns as well as hybridization characteristics and DNA base compositions. The DNA hybridization (S1 nuclease method) revealed five DNA genomic groups, and these groups formed three genomic Nitrobacter species. Mobarry et al. (1996) prepared a hierarchical set of five 16S rRNA-targeted DNA probes for phylogenetically defined groups of autotrophic ammonia- and nitrite-oxidizing bacteria. Their environmental application was demonstrated by quantitative slot blot hybridization and wholecell hybridization of nitrifying activated sludge and biofilm samples. In situ hybridization experiments revealed that Nitrobacter and Nitrosomonas species occurred in clusters and frequently were in contact with each other within sludge flocs. The nitrite oxidoreductase (NOR) from the facultative nitrite-oxidizing bacterium Nitrobacter hamburgensis X14 was used to develop a probe for the gene norB (Kirstein and Bock, 1993). Sequence analysis of DNA fragments revealed three adjacent open reading frames in the order norA, norX and norB. The deduced amino acid sequence of protein NorB contained four cysteine clusters with striking homology to those of iron-sulfur centers of bacterial ferredoxins. Protein NorB shares significant sequence similarity to the β-subunits (NarH and NarY) of the two dissimilatory nitrate reductases (NRA and NRZ) of Escherichia coli. Additionally, the derived amino acid sequence of the truncated open reading frame of norA showed striking resemblance to the α-subunits (NarG and NarZ) of the E. coli nitrate reductases. Additionally, the derived amino acid sequence of the truncated open reading frame of norA showed resemblance to the α-subunits of the E. coli nitrate reductases. Monoclonal antibodies prepared against nitrite oxidoreductases of nitrite oxidizers also

CHAPTER 3.2.17

were used successfully for the identification of nitrite oxidizers (Bartosch et al., 1999). Immunoblot analyses performed with three monoclonal antibodies (MAbs) that recognized the nitrite oxidoreductase (NOR) of the genus Nitrobacter were used for taxonomic investigations of nitrite oxidizers. These MAbs were able to detect the nitrite-oxidizing systems (NOS) of the genera Nitrospira, Nitrococcus and Nitrospina. When the genus-specific reactions of the MAbs were correlated with 16S rDNA sequences, they reflected the phylogenetic relationships among the nitrite oxidizers. In ecological studies, the immunoblot analyses demonstrated that Nitrobacter or Nitrospira cells could be enriched from activated sludge by using various substrate concentrations. The microbiology of the biomass from a nitrite-oxidizing sequencing batch reactor (NOSBR) as well as the seed sludge used were investigated by microscopy, by culturedependent methods, and by molecular biological methods (Burrell et al., 1998). The NOSBR was fed with an inorganic salts solution and nitrite as the sole energy source. It was operated for 6 months, and 16S ribosomal DNA clone libraries were prepared both from the seed sludge and from the reactor. Analysis of the seed sludge revealed that it contained three clones (4% of biomass) that were closely related to the autotrophic nitrite-oxidizer Nitrospira moscoviensis, whereas the NOSBR sludge itself was overwhelmingly dominated by bacteria closely related to the N. moscoviensis (89%). Only two clone sequences were similar to those of the genus Nitrobacter. Near-complete insert sequences of eight clones of N. moscoviensis isolated from the NOSBR and one clone from the seed sludge were determined and phylogenetically analyzed. This report was the first to show the presence of bacteria from the Nitrospira genus in wastewater treatment systems. Nitrite-oxidizing bacteria belonging exclusively to the genus Nitrospira also dominated the nitrite-oxidizing community of a phosphateremoving biofilm from a sequencing batch biofilm reactor (Gieseke et al., 2001). In another study (Burrell et al., 1999), a sequencing batch reactor (SBR) was operated to selectively grow a nitrite-oxidizing microbial community, and it was found that the nitrite oxidation was due the presence of bacteria from the Nitrospira genus and not the Nitrobacter genus, which were in very low numbers. It was hypothesized that the unknown nitrite-oxidizing bacteria in wastewater treatment plants are a range of species related to Nitrospira moscoviensis. Oxidation of nitrite to nitrate in aquaria is typically attributed to bacteria belonging to the genus Nitrobacter that are members of the α-

The Nitrite-Oxidizing Bacteria

865

subdivision of the class Proteobacteria. To identify bacteria responsible for nitrite oxidation in aquaria, clone libraries of rRNA genes were developed by Hovanec et al. (1998) from biofilms from several freshwater aquaria, and analysis of the rDNA libraries, along with results from denaturing gradient gel electrophoresis (DGGE) on frequently sampled biofilms, indicated the presence of putative nitrite-oxidizing bacteria closely related to other members of the genus Nitrospira. Hybridization experiments with rRNA from biofilms of freshwater aquaria demonstrated that Nitrospira-like rRNA comprised nearly 5% of the rRNA extracted from the biofilms during the establishment of nitrification. Nitrite-oxidizing bacteria belonging to the α-subdivision of the class Proteobacteria (e.g., Nitrobacter spp.) were not detected in these samples. Aquaria that received a commercial preparation containing Nitrobacter species did not show evidence of Nitrobacter growth and development but did develop substantial populations of Nitrospira-like species. Time series analysis of rDNA phylotypes on aquaria biofilms by DGGE, combined with nitrite and nitrate analysis, showed a correspondence between the appearance of Nitrospira-like bacterial ribosomal DNA and the initiation of nitrite oxidation. In total, the data suggest that Nitrobacter winogradsky and close relatives were not the dominant nitrite-oxidizing bacteria in freshwater aquaria. Instead, nitrite oxidation in freshwater aquaria appeared to be mediated by bacteria closely related to Nitrospira sp.

Isolation and Nutritional Requirements The information in this section is compiled from the chapter by Bock and Koops (1992) on Nitrobacter in the second edition published in the previous edition of {The Prokaryotes}. Obligate and facultative lithoautotrophic nitrite oxidizers can be successfully isolated only if mineral nitrite media free of any organic contaminants are used, otherwise heterotrophs will overgrow nitrite oxidizers. A sample of 1–2 g (soil, stone, mud etc.) is placed in 100 ml of enrichment medium selective for the growth of either terrestrial or marine strains (Table 1). For the enrichment of dominant nitrifiers, a combination of the most probable number (MPN) technique followed by the serial dilutions method is suitable. In the highest dilutions, nitrite oxidation can be detected only after several months because of the slow growth of the nitrite oxidizers. Plating samples from enrichment cultures on rich and poor solid agar media

866

A. Abeliovich

CHAPTER 3.2.17

Table 1. Four culture mediaa for lithoautotrophic and mixotrophic growth of nitrite oxidizers. Ingredient Distilled water (ml) Seawater (ml) NaNO2 (mg) KNO2 (mg) MgSO4 ·7H2O (mg) CaCl2 ·2H2O (mg) CaCO3 (mg) KH2PO4 (mg) K2HPO4 (mg) FeSO4 (mg) Chelated ironb (mg) Na2Mo4 (µg) (NH4)6Mo7O24 ·4H2O (µg) MnCl2 ·4H2O (µg) CoCl2 ·6H2O (µg) CuSO4 ·5H2O (µg) ZnSO4 ·7H2O (µg) KHCO3 (mg) NaCl (mg) Sodium pyruvate (mg) Yeast extractc (mg) Peptonec (mg)

1a

2b

3b

1,000

1,000

1,000

2,000

2,000

50

50

3.0 150

3.0 150

300 187.5 12.5 500 500 10

4c 300 700 69 100 6.0

1.74 0.15

0.15 1.0 30

50

50 66 0.6 6.0 30

1,500 500

500 550 1,500 1,500

a

pH is adjusted to 7.4–7.8. 13%, Geigy. c Difco. b

can check contamination by heterotrophs. It may take a year to obtain pure cultures with the technique described above. Medium 1a for terrestrial strains is from Aleem and Alexander (1958), media 2b and 3b also for terrestrial strains are from Bock and Stackebrandt (1983), and medium 4c for marine strains is from Watson and Waterbury (1971).

Habitats As described earlier, the nitrite-oxidizing bacteria are ubiquitous, and are usually found in association with the ammonia-oxidizing bacteria, which provide them with their energy source. In spite of many reports describing their sensitivity to light (Guerrero and Jones, 1996; Guerrero and Jones, 1997) and various toxicants (Deni and Penninckx, 1999; Jenicek et al., 1996), nitriteoxidizing bacteria are abundant in practically all terrestrial, marine and fresh water habitats. Nitrifying bacteria play a significant role in erosion of rocks (Lebedeva et al., 1978) primarily because they release nitrates, which dissolve carbonates, in exchange for CO2 they assimilate. This process was found to be a major cause of deterioration of historic monuments in Europe (Mansch and Bock, 1998). Proliferation of nitri-

fiers in sewage is also the basis for nitrogen removal from domestic and industrial waste in water treatment plants, including chemical industry wastewater. Nitrite oxidizers grow within a wide range of pH values up to 10 (Sorokin, 1998a; Sorokin et al., 1998b) and salinities (limits are difficult to determine as nitrifying bacteria resist very low water activities, such as occur when living within desiccated rocks). Although nitrite oxidizers are considered obligate aerobes (even though many can utilize nitrate in absence of oxygen as an electron acceptor), growth of nitrite oxidizers can be observed in totally anaerobic environments such as deep in the mud at the bottom of deep anaerobic wastewater reservoirs, in the absence of nitrate or oxygen, in presence of significant concentrations of sulfides (Abeliovich, 1987). Also, nitrifying bacteria were isolated from a marine anaerobic sediment at depths of up to 8 cm (Blackburn, 1983). Nothing is at present known about the metabolism and survival mechanisms of these nitrite-oxidizing bacteria living under strict anaerobic, reducing conditions. Light (particularly blue light) is frequently mentioned as an inhibiting factor for the nitrification process. However, the reports on the effects of light are sometimes contradictory. Nitrifiers are found in all habitats exposed to sunlight, either at the surface of fresh water or marine habitats, so obviously they must have efficient protection mechanisms to overcome photoinhibition and damage caused by direct sunlight. Sorokin et al. (1998b) and Sorokin (1998a) have isolated five strains of lithotrophic, nitriteoxidizing bacteria from sediments of three soda lakes after enrichment at pH 10 with nitrite as sole energy source; these strains were described as a new species of the genus Nitrobacter, N. alkalicus. Nitrite oxidation to nitrate occurred at pH 10. The nitrifiers had pear-shaped budding cells morphologically similar to those of Nitrobacter and formed tiny colonies on mineral nitrite agar at pH 10. In a study aimed at evaluating the potential of wastewater treatment plants to contaminate receiving waterways, survival of Nitrobacter cells associated with particles in water treatment plant discharges was studied using immunofluoresence methods (Bonnet et al., 1997). It was found that Nitrobacter colonies can settle in freshwater sediments in a week (106 cells per gram of dry sediment) and can therefore colonize river sediments. Chemolithotrophic nitrifying bacteria are dependent on the presence of oxygen for the oxidation of ammonium via nitrite to nitrate. The success of nitrification in oxygen-limited environments largely depends on the oxygen sequestering abilities of both ammonium- and

CHAPTER 3.2.17

nitrite-oxidizing bacteria. Oxygen consumption kinetics were determined with cells grown in mixed culture in chemostats at different growth rates and oxygen tensions (Laanbroek and Gerards, 1993). Reduction of oxygen tension in the culture was found to repress the oxidation of nitrite before the oxidation of ammonium was affected, and hence nitrite accumulated. In addition to the competition between ammonium- and nitrite-oxidizing bacteria, there is competition with organotrophic bacteria for the available oxygen as well. The outcome of the competition is determined by their specific affinities for oxygen as well as by their population sizes. The effect of mixotrophic growth of nitriteoxidizing Nitrobacter hamburgensis on the competition for limiting amounts of oxygen was studied in mixed continuous culture experiments with the ammonium-oxidizing Nitrosomonas europaea at different oxygen concentrations (Laanbroek et al., 1994). The specific affinity for oxygen of N. europaea was in general higher than that of N. hamburgensis, and in transient state experiments, when oxic conditions were switched to anoxic, N. hamburgensis was washed out and nitrite accumulated. However, at low oxygen concentration, the specific affinity for oxygen of N. hamburgensis increased and became as great as that of N. europaea, and owing to its larger population size, the nitriteoxidizing bacterium became the better competitor for oxygen and ammonium accumulated. Therefore Laanbroek et al. suggest that continuously oxygen-limited environments present a suitable ecological niche for the nitrite-oxidizing N. hamburgensis.

Biofilms Biofilms present a unique biotope for using molecular DNA techniques as well as FISH and various sensors. The combination of microsensor and molecular techniques is highly useful for studies on the microbial ecology of biofilms in general, and in particular for the identification of activity sites of nitrifying bacteria. Thus, the distribution of nitrifying bacteria of the genera Nitrosomonas, Nitrosospira, Nitrobacter and Nitrospina was studied by Schramm et al. (2000) in a membrane-bound biofilm system in which gradients of oxygen, pH, nitrite and nitrate were determined by means of microsensors, while the nitrifying populations along these gradients were identified and quantified using FISH in combination with confocal laser scanning microscopy. It was found that the oxic part of the biofilm was dominated by ammonia oxidizers and by members of the genus Nitrobacter, and Nitrospira sp. were virtually absent in this

The Nitrite-Oxidizing Bacteria

867

part of the biofilm, whereas they were most abundant at the oxic-anoxic interface. In the totally anoxic part of the biofilm, cell numbers of all nitrifiers were relatively low. These observations suggest the microaerophilic behavior of an as yet uncultured Nitrospira sp. as a factor affecting its environmental competitiveness. De Beer and Schramm (1999), using microsensors and molecular techniques (such as in situ hybridization with 16S rDNA-targeted oligonucleotide probes), showed that there exists in biofilms grown in bioreactors a complex nitrifying community, consisting of members of the genera Nitrosomonas, Nitrosospira, Nitrobacter and Nitrospira. In nitrifying biofilms from a trickling filter of an aquaculture water recirculation system, it was found that nitrification was restricted to a narrow zone of 50 µm of the very top of the film. Ammonia oxidizers formed a dense layer of cell clusters in the upper part of the biofilm, whereas the nitrite oxidizers showed less-dense aggregates in the proximity of the ammonia oxidizers. Both ammonia and nitrite oxidizers were not restricted to the oxic zone of the biofilm but were also detected in substantially lower numbers in the anoxic layers and even occasionally at the bottom of the biofilm (Schramm et al., 1996).

Enumeration Conventional methodologies for counting nitrifiers are gradually being replaced by new technologies, although the MPN method still dominates, this because all methods based on specific probes or antibodies still leave open the possibility that some species do not crossreact with the probe used. On the other hand, the issue of optimal conditions for maximal yields using MPN counts has not been yet resolved. One example is a new amperometric enzymelinked immunoassay for specific enumeration of Nitrobacter, which uses an electrode made of glassy carbon on which an immunological reaction is carried out. The detection limit was approximately 3 × 106 Nitrobacter cells/ml (Sanden and Dalhammar, 2000). It was shown that this method could be applied for the enumeration of Nitrobacter in activated sludge and other environmental samples. The reason for the attempts to develop new methodologies is the lengthy incubation periods required for stable MPN counts: maximum most probable numbers of the ammonia-oxidizing group were attained in 20–55 days (median, 25) and MPN estimates of nitrite oxidizers required a much longer incubation (103–113 days; Matulewich et al., 1975). In addition, Both et al. (1990a) and (1990b) studied the effect of two concentrations of nitrite in the

868

A. Abeliovich

incubation medium, 0.05 and 5.0 mM, and found that numbers of nitrite-oxidizing bacteria were highly dependent on the nitrite concentration as well as on the soil sampled. Neither the influence of pH or nitrite concentration in the incubation medium could account exclusively for the MPN-enumeration result, which raises the issue of whether to use more than one incubation medium for the enumeration of nitrite-oxidizing bacteria. Enumeration of nitrite-oxidizing bacteria in soil samples by the MPN technique often showed relatively high cell number at a low nitrite concentration, when compared to the number of ammoniumoxidizing bacteria. When different Nitrobacter species as well as nongrowing cells differing in age were incubated 5 months at 20°C in presence of various nitrite concentrations and pH values, it appeared that the growth of cells taken from an early stationary phase culture of all these species was not affected by high nitrite concentrations or low pH. Growth of 8- and 18-month-old nongrowing cells of Nitrobacter hamburgensis was also insensitive to high nitrite concentration (5 mM). The growth of 8- and 18-month old resting cells of N. vulgaris was repressed only by a combination of 5 mM nitrite and a low pH. Growth of 8-month-old nongrowing cells of N. winogradsky was sensitive to 5 mM irrespective of pH, but growth of 18-month-old cells were inhibited by 5 mM nitrite only at a low pH. The growth of 8- and 18-month-old resting cells of N. winogradsky serotype agilis was repressed by low pH rather than by high nitrite concentration (Laanbroek and Schotman, 1991). These results emphasize the problems associated with using MPN counts for estimating size of nitrifier populations in natural samples. A molecular approach based on PCR that was expected to detect and quantify nitrifying bacteria was also tested using specific primers of the genus Nitrobacter. In this study, coupled in parallel was a 14C-radiotracing method used to measure potential nitrification; it was shown that Nitrobacter represented less than 0.1% of the total bacterial community (Berthe et al., 1999). In another study, also aimed at counting Nitrobacter populations in situ by PCR, two primers from the 16S rDNA gene were used to generate a 397-bp fragment by amplification of Nitrobacter species DNA, and it was found that the PCR had a lower detection threshold (102 Nitrobacter cells per gram of soil) than did the MPN or fluorescent antibody method (Degrange and Bardin, 1995). In contrast, another study (Feray et al., 1999) found the best recovery yield was obtained with the immunofluorescence technique (21.3%), and the poorest detection level was reached with the MPN method (3.1%).

CHAPTER 3.2.17

Wastewater Tertiary wastewater treatment, aimed at removing all nitrogen wastes from the effluents, is costly in terms of the oxygen required for oxidizing ammonia to nitrate. However, as far as the nitrification—denitrification process is concerned, oxidation of nitrite to nitrate is an unnecessary step, as denitrification can just as well proceed from nitrite. Therefore, attempts are being made to devise operational parameters that will inhibit the activity of nitrite-oxidizing bacteria in wastewater treatment plants. Thus, it was found (Rhee et al., 1997) that accumulation of nitrite occurred during the aerobic phase of a sequencing batch reactor (SBR) operating to remove nitrogen from synthetic wastewater: the activity of autotrophic nitrite oxidizers was reduced in the SBR and the free ammonia was the main inhibitor of nitrite oxidation. It has been shown experimentally (Garrido et al., 1997) that it is possible to convert all ammonium to approximately 50% nitrate and 50% nitrite in the effluent of a biofilm air-lift suspension reactor, this with oxygen concentrations between 1 and 2 mg/liter. The ammonia- and nitrite-oxidizing bacterial populations occurring in the nitrifying activated sludge of an industrial plant treating sewage with high ammonia concentrations were also studied using in situ hybridization with a set of hierarchical 16S rDNA-targeted probes (Juretschko et al., 1998). Although a Nitrobacter strain was isolated, members of the genus Nitrobacter were not detectable in the activated sludge by in situ hybridization. A specific 16S rDNA-targeted probe for Nitrospira demonstrated that a Nitrospira-like bacterium was present in significant numbers (9% of the total bacterial counts). According to Blackall (2000), even though it is common knowledge that Nitrosomonas and Nitrobacter are the major ammonia and nitrite oxidizers, today we know that these organisms may not play any role in the transformations for which they have achieved such acclaim. Using the above-mentioned methodologies, Nitrospiralike bacteria were found to be the dominant nitrite oxidizers in both enriched and full scale nitrifying systems.

Resistance, Inhibition and Biodegradation Because inhibition of nitrite oxidation is economically advantageous in wastewater treatment plants, procedures were developed for identification of nitrification inhibitors in wastewater.

CHAPTER 3.2.17

These include fractionation of wastewater samples and a nitrification inhibition assay with pure cultures of Nitrobacter to identify the inhibitory effect. In such a study, a series of unsaturated fatty acids and two monoterpenes were found to be inhibitory in an industrial wastewater sample (Svenson et al., 2000). A shorter nitrification-denitrification cycle occurs in the presence of free ammonium. Although the occurrence of this cycle depends on both a high ammonia concentration and high pH, it was found that the pH of the wastewater is the decisive parameter (Surmacz-Gorska et al., 1997). However, a study aimed at identifying nitrification at high pH values in soda lakes and soda soils with pH 9.5–11 led to the isolation of a nitrifier morphologically similar to Nitrobacter that formed tiny colonies on mineral nitrite agar at pH 10 (Sorokin, 1998a). Hwang et al. (2000), in a detailed study of the inverse relation between alkalinity and ammonia with respect to inhibition of nitrite oxidation, showed that when the molar ratio of carbonate alkalinity to ammonia increased from 4.1 to 9.4 (thus increasing the concentration of free ammonia), the ammonium removal rate doubled. At the same time the higher concentration of free ammonia in the medium was a selective inhibitor for Nitrobacter, causing an enhanced nitrite build-up in a biofilm reactor. As for the effect of heavy metals, laboratory evaluations were conducted to study the toxic effects of copper and nickel on a culture of strictly obligatory nitrifiers (Nitrosomonas sp. and Nitrobacter sp.) in continuous flow stirred tank reactors. Nitrosomonas sp. was found to be equally or more sensitive than Nitrobacter sp. (Lee et al., 1997). Attempts were made to develop a process for the simultaneous removal of organic halogens and nitrogen from kraft pulp mill effluents. A nitrifying biofilm reactor removed organic halogens from bleached kraft pulp mill effluents including chlorophenols from synthetic wastewater (Kostyal et al., 1997; Kostyal et al., 1998). In another study, none of several chemicals tested (nonylphenol, naphthalene, 2-methylnaphthalene, di-2-ethylhexylphthalate and toluene) inhibited nitrification when added to soil at various concentrations (Kirchmann et al., 1991). However, in many cases municipal wastewater did contain substances that inhibited nitrification to varying degrees (Jonsson et al., 2000). Reports conflict on the effect of toluene on indigenous microbial populations. Nitrite oxidation potential (NOP) was reduced after incubation with high toluene concentrations for 45 days (Fuller and Scow, 1996). Trichloroethylene is also inhibitory to the soil indigenous nitrifying population (Fuller and Scow, 1997).

The Nitrite-Oxidizing Bacteria

869

Volatile fatty acids such as formic, acetic, propionic, n-butyric, isobutyric, n-valeric, isovaleric and n-caproic acid as well as trimethylamine are inhibitory to nitrification, but only at high concentrations, suggesting that volatile fatty acids and trimethylamine alone cannot account for the inhibition of the nitrification activity in domestic wastewater (Eilersen et al., 1994). Nitrifying bacteria were also found to be resistant to monochloramines, and the combination of increased concentrations of oxidized nitrogen with decreased total chlorine in treated water was used as an indicator of bacterial nitrification (Cunliffe, 1991). The need to set limits on nitrite oxidation in wastewater treatment plants created a demand for assays to determine the degree of inhibition of nitrification. One such assay, developed by Grunditz and Dalhammar, was based on inhibition of pure cultures of Nitrosomonas and Nitrobacter isolated from sewage sludge (Grunditz and Dalhammar, 2001). As for the effects of light, nitrifying bacteria (both ammonia and nitrite oxidizers) are capable of recovery from photoinhibition in the dark (see above). Recovery of oxidizing activities is both dose and wavelength dependent (Guerrero and Jones, 1996). The light-absorbing pigment was identified as a porphyrin-like pigment with an absorption maximum at 408 nm, accumulating at the late exponential phase of growth. This photoreceptor was found at higher concentrations in ammonia oxidizers than in nitrite oxidizers (Guerrero and Jones, 1997).

Lithotrophy An evaluation of field data from historical buildings in Germany, carried on by Mansch and Bock (1998), showed that ammonia and nitrite oxidizers were found in 55 and 62% of the samples, respectively, and that nitrifying bacteria will colonize natural stone within several years of exposure. The highest cell numbers were in some cases found underneath the surface. Nitrifying bacteria showed a preference for calcareous material with a pore radius of 1–10 µm. Cell numbers of nitrifying bacteria did not correlate with the nitrate content of the stone material. Their data strongly suggest that microbial colonization of historical buildings is enhanced by anthropogenic air pollution. A comparison of samples taken between 1990 and 1995 from buildings throughout Germany showed that in eastern Germany significantly more colonization with facultatively methylotrophic bacteria and nitrifying bacteria existed. The same was true for natural stone from an urban exposure site when compared to material from a rural exposure site.

870

A. Abeliovich

CHAPTER 3.2.17

Literature Cited

Eilersen, A. M., M. Henze, and L. Kloft. 1994. Effect of volatile fatty-acids and trimethylamine On nitrification in activated-sludge. Water Res. 28:1329–1336. Feray, C., B. Volat, V. Degrange, A. Clays-Josserand, and B. Montuelle. 1999. Assessment of three methods for detection and quantification of nitrite-oxidizing bacteria and nitrobacter in freshwater sediments. Microb. Ecol. 37:208–217. Fuller, M. E., and K. M. Scow. 1996. Effects of toluene on microbially-mediated processes involved in the soil nitrogen cycle. Microb. Ecol. 32:171–184. Fuller, M. E., and K. M. Scow. 1997. Impact of trichloroethylene and toluene on nitrogen cycling in soil. Appl. Environ. Microbiol. 63:4015–4019. Garrido, J. M., W. A. J. van Benthum, M. C. M. van Loosdrecht, and J. J. Heijnen. 1997. Influence of dissolved oxygen concentration on nitrite accumulation in a biofilm airlift suspension reactor. Biotechnol. Bioeng. 53:168–178. Gieseke, A., U. Purkhold, M. Wagner, R. Amann, and A. Schramm. 2001. Community structure and activity dynamics of nitrifying bacteria in a phosphate-removing biofilm. Appl. Environ. Microbiol. 67:1351–1362. Grunditz, C., and G. Dalhammar. 2001. Development of nitrification inhibition assays using pure cultures of Nitrosomonas and Nitrobacter. Water Res. 35:433–440. Guerrero, M. A., R. D. Jones. 1996. Photoinhibition of marine nitrifying bacteria. 2: Dark recovery after monochromatic or polychromatic irradiation. Marine Ecology (Progress Series) 141:193–198. Guerrero, M. A., and R. D. Jones. 1997. Light-induced absorbance changes associated with photoinhibition and pigments in nitrifying bacteria. Aquat. Microb. Ecol. 13:233–239. Guschin, D. Y., B. K. Mobarry, D. Proudnikov, D. A. Stahl, B. E. Rittmann, and A. D. Mirzabekov. 1997. Oligonucleotide microchips as genosensors for determinative and environmental studies in microbiology. Appl. Environ. Microbiol. 63:2397–2402. Hovanec, T. A., L. T. Taylor, A. Blakis, and E. F. Delong. 1998. Nitrospira-like bacteria associated with nitrite oxidation in freshwater aquaria. Appl. Environ. Microbiol. 64: 258–264. Hwang, B. H., K. Y. Hwang, E. S. Choi, D. K. Choi, and J. Y. Jung. 2000. Enhanced nitrite build-up in proportion to increasing alklinity/NH4+ ratio of influent in biofilm reactor. Biotchnol. Lett. 22:1287–1290. Jenicek, P., J. Zabranska, and M. Dohanyos. 1996. The influence of anaerobic pretreatment on the nitrogen removal from biosynthetic pharmaceutical wastewaters. Ant. v. Leeuwenhoek 69:41–46. Jonsson, K., C. Grunditz, G. Dalhammar, and J. L. Jansen. 2000. Occurrence of nitrification inhibition in Swedish municipal wastewaters. Water Res. 34:2455–2462. Juretschko, S., G. Timmermann, M. Schmid, K. H. Schleifer, A. Pommerening-Roser, H. P. Koops, and M. Wagner. 1998. Combined molecular and conventional analyses of nitrifying bacterium diversity in activated sludge: Nitrosococcus mobilis and Nitrospira-like bacteria as dominant populations. Appl. Environ. Microbiol. 64:3042–3051. Kirchmann, H., H. Astrom, and G. Jonsall. 1991. Organic pollutants in sewage-sludge. 1: Effect of toluene, naphthale, 2-methylnaphthalene, 4-normal-nonylphenol and di-2-ethylhexyl phthalate on soil biological processes and their decomposition in soil. Swedish J. Agric. Res. 21:107–113.

Abeliovich, A. 1987. Nitrifying bacteria in wastewater reservoirs. Appl. Environ. Microbiol. 53:754–760. Aleem, M. I. H., and M. Alexander. 1958. Cell free nitrification by nitrobacter. J. Bacteriol. 76:510–514. Bartosch, S., I. Wolgast, E. Spieck, and E. Bock. 1999. Identification of nitrite-oxidizing bacteria with monoclonal antibodies recognizing the nitrite oxidoreductase. Appl. Environ. Microbiol. 65:4126–4133. Berthe, T., J. Garnier, and F. Petit. 1999. Quantification of nitrifying bacteria of the genus Nitrobacter in an aquatic system (Seine estuary, France). Science 322:517–526. Blackall, L. L. 2000. A summary of recent microbial discoveries in biological nutrient removal from wastewater. Austral. Biotechnol. 10:29–31. Blackburn, H. 1983. The microbial nitrogen cycle. In: W. E. Krumbein (Ed.) Microbial Geochemistry. Blackwell Scientific Publications. Boston, MA. 63–89. Bock, E., H. Sundermeyer-Klinger, and E. Stackebrandt. 1983. New facultative lithoautotrophic nitrite oxidizing bacteria. Arch. Microbiol. 136:281–284. Bock, E., and H. P. Koops. 1992. The genus nitrobacter and related genera. In: A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K. H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York, NY. 3:2302–2309. Bonnet, C., B. Volat, R. Bardin, V. Degranges, and B. Montuelle. 1997. Use of immunofluorescence technique for studying a Nitrobacter population from wastewater treatment plant following discharge in river sediments: First experimental data. Water Res. 31:661–664. Both, G. J., S. Gerards, and H. J. Laanbroek. 1990a. Enumeration of nitrite oxidizing bacteria in grassland soils using a most probable technique: Effect of nitrite concentration and sampling procedure. FEMS Microbiol. Ecol. 74:277–285. Both, G. J., S. Gerards, and H. J. Laanbroek. 1990b. Most probable numbers of chemlitho-autotrophic nitriteoxidizing bacteria in well drained grassland soils: Stimulation by high nitrite concentrations. FEMS Microbiol. Ecol. 74:287–293. Burrell, P. C., J. Keller, and L. L. Blackall. 1998. Microbiology of a nitrite-oxidizing bioreactor. Appl. Environ. Microbiol. 64:1878–1883. Burrell, P., J. Keller, and L. L. Blackall. 1999. Characterisation of the bacterial consortium involved in nitrite oxidation in activated sludge. Water Sci. Technol. 39:45–52. Cunliffe, D. A. 1991. Bacterial nitrification in chloraminated water supplies. Appl. Environ. Microbiol. 57:3399– 3402. de Beer, D., and A. Schramm. 1999. Micro-environments and mass transfer phenomena in biofilms studied with microsensors. Water Sci. Technol. 39:173–178. Degrange, V., and R. Bardin. 1995. Detection and counting of Nitrobacter populations in soil by PCR. Appl. Environ. Microbiol. 61:2093–2098. Deni, J., and M. J. Penninckx. 1999. Nitrification and autotrophic nitrifying bacteria in a hydrocarbonpolluted soil. Appl. Environ. Microbiol. 65:4008–4013. Ehrich, S., D. Behrens, E. Lebedeva, W. Ludwig, and E. Bock. 1995. A new obligately chemolithoautotrophic, nitriteoxidizing bacterium, Nitrospira moscoviensis sp. nov. and its phylogenetic relationship. Arch. Microbiol. 164:16–23.

CHAPTER 3.2.17 Kirstein, K., and E. Bock. 1993. Close genetic relationship between Nitrobacter hamburgensis nitrite oxidoreductase and Escherichia coli nitrate reductases. Arch. Microbiol. 160:447–453. Kostyal, E., E. L. Nurmiaho Lassila, J. A. Puhakka, and M. Salkinoja Salonen. 1997. Nitrification, denitrification, and dechlorination in bleached kraft pulp mill wastewater. Appl. Microbiol. Biotechnol. 47:734–741. Kostyal, E., M. Borsanyi, L. Rigottier-Gois, and M. S. Salkinoja-Salonen. 1998. Organic halogen removal from chlorinated humic ground water and lake water by nitrifying fluidized-bed biomass characterised by electron microscopy and molecular methods. Appl. Microbiol. Biotechnol. 50:612–622. Laanbroek, H. J., and J. M. T. Schotman. 1991. Effect of nitrite concentration and pH on most probable number enumerations of non growing nitrobacter spp. FEMS Microbiol. Ecol. 85:269–278. Laanbroek, H. J., and S. Gerards. 1993. Competition for limiting amounts of oxygen between Nitrosomonas europaea and Nitrobacter winogradskyi grown in mixed continuous cultures. Arch. Microbiol. 159:453–459. Laanbroek, H. J., P. L. E. Bodelier, and S. Gerards. 1994. Oxygen consumption kinetics of Nitrosomonas europaea and Nitrobacter hamburgensis in mixed continuous cultures at different oxygen concentrations. Arch. Microbiol. 161:156–162. Lebedeva E. V., N. N. Lialikova, and I. Bugel’skii. 1978. Participation of nitrifying bacteria in the disintegration of serpentinous ultrabasic rock. Mikrobiologiia 47:1101– 1107. Lee, Y. W., S. K. Ong, and C. Sato. 1997. Effects of heavy metals on nitrifying bacteria. Water Sci. Technol. 36:69– 74. Macdonald, R. M. 1986. Nitrification in soil: An introductory history. In: J. I. Prosser (Ed.) Nitrification. IRL Press. Washington, DC, Washington, DC. 20:1–17. Mansch, R., and E. Bock. 1998. Biodeterioration of natural stone with special reference to nitrifying bacteria. Biodegradation 9:47–64. Matulewich, V. A., P. F. Strom, and M. S. Finstein. 1975. Length of incubation for enumerating nitrifying bacteria present in various environments. Appl. Microbiol. 29:265–268. Meincke, M., E. Bock, D. Kastrau, and P. M. H. Kroneck. 1992. Nitrite oxidoreductase from Nitrobacterhamburgensis-redox centers and their catalytic role. Arch. Microbiol. 158:127–131. Mobarry, B. K., M. Wagner, V. Urbain, B. E. Rittmann, and D. A. Stahl. 1996. Phylogenetic probes for analyzing abundance and spatial organization of nitrifying bacteria. Appl. Environ. Microbiol. 62:2156–2162. Navarro, E., M. P. Fernandez, F. Grimont, A. Claysjosserand, and R. Bardin. 1992a. Genomic heterogeneity of the genus Nitrobacter. Int. J. Syst. Bacteriol. 42:554–560. Navarro, E., P. Simonet, P. Normand, and R. Bardin. 1992b. Characterization of natural populations of Nitrobacter spp. using PCR/RFLP analysis of the ribosomal intergenic spacer. Arch. Microbiol. 157:107–115. Noda, N., H. Ikuta, Y. Ebie, A. Hirata, S. Tsuneda, M. Matsumura, T. Sumino, and Y. Inamori. 2000. Rapid quantification and in situ detection of nitrifying bacteria in biofilms by monoclonal antibody method. Water Sci. Technol. 41:301–308. Okabe, S., H. Satoh, and Y. Watanabe. 1999. In situ analysis of nitrifying biofilms as determined by in situ hybridiza-

The Nitrite-Oxidizing Bacteria

871

tion and the use of microelectrodes. Appl. Environ. Microbiol. 65:3182–3191. Orso, S., M. Gouy, E. Navarro, and P. Normand. 1994. Molecular phylogenetic analysis of Nitrobacter spp. Int. J. Syst. Bacteriol. 44:83–86. Pasteur, L. 1862. Etudes sur les mycoderms: Rôle de ces plantes dans la fermentation acétique. C. R. Acad. Sci. 54:265–270. Rhee, S. K., J. J. Lee, and S. T. Lee. 1997. Nitrite accumulation in a sequencing batch reactor during the aerobic phase of biological nitrogen removal. Biotechnol. Lett. 19:195– 198. Sakai, K., Y. Ikehata, Y. Ikenaga, M. Wakayama, and M. Moriguchi. 1996. Nitrite oxidation by heterotrophic bacteria under various nutritional and aerobic conditions. J. Ferment. Bioeng. 82:613–617. Sakai, K., K. Nakamura, M. Wakayama, and M. Moriguchi. 1997. Change in nitrite conversion direction from oxidation to reduction in heterotrophic bacteria depending on the aeration conditions. J. Ferment. Bioeng. 84:47–52. Sanden B., and G. Dalhammar. 2000. Application of an amperometric immuno sensor for the enumeration of Nitrobacter in activated sludge. Appl. Microbiol. Biotechnol. 54:413–417. Schramm, A., L. H. Larsen, N. P. Revsbech, N. B. Ramsing, R. Amann, and K. H. Schleifer. 1996. Structure and function of a nitrifying biofilm as determined by in situ hybridization and the use of microelectrodes. Appl. Environ. Microbiol. 62:4641–4647. Schramm, A., D. De Beer, M. Wagner, and R. Amann. 1998. Identification and activities in situ of Nitrosospira and Nitrospira spp. as dominant populations in a nitrifying fluidized bed reactor. Appl. Environ. Microbiol. 64: 3480–3485. Schramm, A., D. De Beer, A. Gieseke, and R. Amann. 2000. Microenvironments and distribution of nitrifying bacteria in a membrane-bound biofilm. Environ. Microbiol. 2:680–686. Sorokin, D. Y. 1991. Peroxide mechanism of nitrite oxidation in heterotrophic bacteria. Microbiology 60:4–9. Sorokin, D. Y. 1998a. On the possibility of nitrification in extremely alkaline soda biotopes. Microbiology 67:335– 339. Sorokin, D. Y., G. Muyzer, T. Brinkhoff, J. G. Kuenen, and M. S. Jetten. 1998b. Isolation and characterization of a novel facultatively alkaliphilic Nitrobacter species, N. alkalicus sp. nov. Arch. Microbiol. 170: 345–352. Surmacz-Gorska, J., A. Cichon, and K. Miksch. 1997. Nitrogen removal from wastewater with high ammonia nitrogen concentration via shorter nitrification and denitrification. Water Sci. Technol. 36:73–78. Svenson, A., B. Sanden, G. Dalhammar, M. Remberger, and L. Kaj. 2000. Toxicity identification and evaluation of nitrification inhibitors in wastewaters. Environ. Toxicol. 15:527–532. Teske, A., E. Alm, J. M. Regan, S. Toze, B. E. Rittmann, and D. A. Stahl. 1994. Evolutionary relationships among ammonia- and nitrite-oxidizing bacteria. J. Bacteriol. 176:6623–6630. Watson, S. W., and J. B. Waterbury. 1971. Characteristics of two marine nitrite oxidizing bacteria, Nitrospina gracilis nov. gen. nov. sp. and Nitrococcus mobilis nov. gen. nov. sp. Arch. Mikrobiol. 77:203–230. Watson, S. W., F. W. Valois, and J. B. Waterbury. 1981. The family Nitrobacteraceae. In: M. P. Starr, H. Stolp, H. G. Trüper, A. Balows, and H. G. Schlegel (Eds.) {The

872

A. Abeliovich

Prokaryotes}. Springer-Verlag. New York, NY. 1:1005– 1022. Watson, S. W., E. Bock, F. W. Valois, J. B. Waterbury, and U. Schlosser. 1986. Nitrospira marina gen. nov., sp. nov.: A chemolithotophic nitrite oxidizing bacterium. Arch. Microbiol. 144:1–7. Woese, C. R., W. G. Weisburg, B. J. Paster, C. M. Hahn, R. S. Tanner, N. R. Krieg, H. P. Koops, H. Harms, and

CHAPTER 3.2.17 E. Stackebrandt. 1984. The phylogeny of purple bacteria: The beta subdivision. Syst. Appl. Microbiol. 5:327–336. Woese, C. R., W. G. Weisburg, C. M. Hahn, B. J. Paster, L. B. Zablen, B. J. Lewis, T. J. Macke, W. Ludwig, and E. Stackebrandt. 1985. The phylogeny of purple bacteria: The gamma subdivision. Syst. Appl. Microbiol. 6:25– 33.

Prokaryotes (2006) 5:873–891 DOI: 10.1007/0-387-30745-1_42

CHAPTER 3.2.18 ehT

areneG

, suc raozA

, o i rb i vozA

ar i psozA

dna

The Genera Azoarcus, Azovibrio, Azospira and Azonexus BARBARA REINHOLD-HUREK AND THOMAS HUREK

Introduction The genera Azoarcus, Azospira, Azovibrio and Azonexus harbor mostly nitrogen-fixing Betaproteobacteria. In their first taxonomic description, several strains which were later assigned to the genera Azovibrio, Azospira (Reinhold-Hurek et al., 1993) and Azonexus (Hurek et al., 1997b) were included in the genus Azoarcus sensu lato. Therefore, they are discussed in one chapter. Except for one strain, all these diazotrophic bacteria had been isolated from a similar source, roots of a pioneer grass in Pakistan or fungal resting stages from the same field; they had many physiological features in common, and they were phylogenetically related. Therefore, three groups of bacteria were included into Azoarcus although they were located on the Azoarcus rRNA branch at low Tm(e) values (Reinhold-Hurek et al., 1993) or showed a low 16S rDNA similarity (Hurek et al., 1997b). Isolation of additional strains, which made possible a better phenotypic description, and sequencing of almost complete 16S rDNA sequences allowed the reassessment of the taxonomic structure of Azoarcus sensu lato. The unnamed groups C and D (Reinhold-Hurek et al., 1993) obtained the rank of different genera Azovibrio and Azospira, respectively (ReinholdHurek and Hurek, 2000); Azoarcus sensu lato group E (Hurek et al., 1997b) was proposed as Azonexus (Hurek et al., 1997b). Strains of these genera deserve special recognition as endophytes of grass roots or as degraders of aromatic compounds.

The Genus Azoarcus Introduction The genus Azoarcus harbors two ecologically different groups of bacteria: 1) soil-borne strains which can degrade aromatic hydrocarbons under denitrifying conditions, and 2) bacteria associated with grass roots epi- or endophytically, which apparently do not survive well in root-free soil. They will be referred to as “soil-borne” or

“plant-associated” species, respectively, in this chapter. Azoarcus was originally described with two valid species of grass-associated diazotrophs (Reinhold-Hurek et al., 1993). However, a growing number of species of soil-borne strains has been added in recent years.

Phylogeny Azoarcus spp. are members of the Betaproteobacteria in the Rhodocyclales according to phylogenetic analysis of almost complete 16S rRNA gene sequences. They are phylogenetically most closely related to Thauera with which they form the Azoarcus/Thauera branch (Fig. 1). Azoarcus species are located on two different clades, which in part reflects their physiology and ecology: the soil-borne species A. tolulyticus, A. toluclasticus, A. toluvorans, A. evansii, A. buckelii and A. anaerobius and the plant-associated species A. indigens, A. communis and Azoarcus sp. strain BH72. The analysis of phylogenetic relationships within the Azoarcus/Thauera 16S rDNA cluster is rendered difficult because the branching pattern between Thauera, the soil-borne and the plant-associated Azoarcus species is unstable when tested with different tree-building methods. This was also observed previously (Hurek et al., 1997b; Reinhold-Hurek and Hurek, 2000). Also in other taxa closely related to each other, like Rhizobium and related genera, the resolution of phylogenetic analysis is sometimes limited. Nevertheless, all named species in the genus Thauera cluster in one clade with highly significant support levels (Fig. 1), allowing a reliable assignment to this genus. Also most soil-borne species or strains of Azoarcus fall into one clade with significant support levels, and none of the species containing plant-associated strains are located on this branch. The 16S rDNA phylogenetic distances within these three clades are similar. Within the genus Azoarcus, they are up to 6%, while Thauera is 6–7% distant. Thus Azoarcus spp. represents a rather heterogeneous group. Moreover, since the phylogenetic distances within the three clades in the Azoarcus/ Thauera group are similar, both subgroups of

suxenozA

874

B. Rareinhold-Hurek and T. Hurek

CHAPTER 3.2.18 Azoarcus sp. Td-19 (L33690) Azoarcus sp. Td-3 (L33693) Azoarcus sp. ToN1 (X83534) 75 Azoarcus evansii pF6. (U44853) Azoarcus evansii KB740 r (X77679) 92 26 Azoarcus toluvorans Td-21 r (L33692) Azoarcus sp. M3 (Y11040) 52 37 Azoarcus sp. Td-15 (L33688) 83 Azoarcus sp. T (AF129465) 89 Azoarcus toluvorans Td-17 (L33689) 44 Azoarcus sp. CC-11 (AB033745) 68 Azoarcus sp. FL05 (AF011330) Azoarcus toluclasticus MF7 (AF123076) 50 Azoarcus toluclasticus MF63 r (AF123077) 96 Azoarcus tolulyticus Td-2 (L33691) 61 84 Azoarcus tolulyticus BL11 (AF123075) 56 Azoarcus tolulyticus Td-1 (L33687) 53 Azoarcus tolulyticus Tol-4 r (L33694) Azoarcus sp. 22Lin (Y13222) Azoarcus sp. T3 (Y11041) Azoarcus sp. CR23 (AF011328) 62 Azoarcus sp. EbN1 (X83531) 99 Azoarcus buckelii U120 (AJ315676) Azoarcus anaerobius LuFRes1 r (Y14701) 74 Azoarcus sp. HxN1 (AF331975) 38 Azoarcus sp. PBN1 (X83532) 88 71

43 44

47 97

55

71

25

52 100 63 100 52 80

96 100

70

50

43 63 44

38

32

48 20 23

Uncultured bacterium clone A33 (AF072925) Uncultured bacterium clone H30 (AF072924) Uncultured bacterium clone H25 (AF072919) 100 Uncultured bacterium clone S23 (AF072921) 85 83 Uncultured bacterium clone S3 (AF072918) 100 Azoarcus indigens ME14 Azoarcus indigens VB32 r (AF011345) Azoarcus sp. BH72 (AF011344) Azoarcus sp. LU1 (AJ007007) Azoarcus communis SWub3 r (AF011343) 95 Azoarcus communis ME91 Uncultured bacterium clone OS ac16 Thauera

Duganella 98 Azovibrio restrictus S5b2 r (AF011346) Azovibrio sp. BS20-3 (AF011349) Uncultured bacterium clone SJA-10 (AJ009452) Azonexus fungiphilus BS5-8 r (AF011350) Dechloromonas sp. MissR (AF170357) 66 48 Dechloromonas sp. SIUL (AF170356) 55 Dechloromonas sp. JJ (AY032611) 72 Ferribacterium limneticum cda-1 (Y17060) 46 Isolate OcN1 (AF331976) Isolate B2 (AF035048) Uncultured bacterium clone SBRH147 (AF204246) 100 25 Dechloromonas agitatus CBK (AF047462) 44 100 Dechloromonas sp. CL (AF170354) Dechloromonas sp. NM (AF170355) 86 Uncultured bacterium clone SA35 (AF245350) Uncultured bacterium clone PHOS-HE23 (AF314420) Isolate B5 (AF035051) 100 Isolate 51885 (AF227840) Oral isolate A08KA (AY005031) Rhodocyclus tenuis DSM 110 (D16209) Rhodocyclus purpureus DSM 168 (M34132) 96 Uncultured bacterium clone SJA-109 (AJ009484) 97 Propionivibrio dicarboxylicus DSM 5885 (Y17601) Propionibacter pelophilus asp 66 (AF016690) Azospira oryzae 6a3 (AF011347) 96 Uncultured bacterium clone SJA-52 (AJ009466) Dechlorosoma] sp. SDGM (AF170349) 100 64 Dechlorosoma] sp. Iso2 (AF170351) 51 Dechlorosoma] sp. Iso1 (AF170350) 48 Dechlorosoma suillum] ps (AF170348)

0.01

Fig. 1. Phylogenetic analysis of 16S rDNA sequences (1358 positions) of a subgroup of the Betaproteobacteria. Subtree of the Rhodocyclus/Thauera/Azoarcus group derived from an analysis of 159 sequences. Tree inference was carried out using the neighbor-joining algorithm and a Jukes-Cantor correction with 125 bootstrap repetitions. Sequence accession numbers are given in parentheses.

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Azoarcus might also deserve the rank of different genera in the future. However, this will require a rigid polyphasic taxonomic analysis. On the basis of 16S rDNA sequence analyses, all of the validly described species of Azoarcus are well resolved except for the two strains of A. toluvorans, which are interdispersed with several strains of uncertain affiliation (Fig. 1). Former members of A. tolulyticus, Td-3 and Td-19, have recently been removed from this species owing to low DNA-DNA similarity (below 40%), which is also reflected in our phylogenetic analysis (Fig. 1). Interestingly, also a deeply-branching clade of as yet uncultured bacteria detected in nitrifyingdenitrifying activated sludge of an industrial wastewater treatment plant (Juretschko et al., 2002) is localized within the Azoarcus cluster (Fig. 1).

875

and Hurek, 2000; see also the section on Identification in this Chapter). Therefore, both subgroups of Azoarcus might deserve the rank of different genera in future. However, this will require a rigid polyphasic taxonomic analysis. As can be seen from the descriptive table (Table 3), many features have not been tested for all species or strains. There appears to be a bias in testing, e.g., nitrogen fixation and aerobic nutritional profiles are not excessively tested in soil-borne strains, whereas nutritional profiles under conditions of denitrification are missing for plantassociated species. These studies should also include the numerous soil-borne isolates not yet assigned to any species. As some strains are deeply branching in the phylogenetic analysis (e.g., 22Lin, T3 and M3; Fig. 1), the description of new species might be expected.

Habitat Taxonomy The definition of the genus Azoarcus is quite complex. It includes strains with a variety of physiological and ecological properties, which place them into at least two groups. In addition to their ecology, the species containing plantassociated strains differ from the soil-borne species in some nutritional features, e.g., in their inability to use carbohydrates as sole carbon source or, under conditions of denitrification, a wide range of aromatic compounds. The genus Azoarcus consists of the following eight validly described species, mostly based on polyphasic taxonomic approaches including DNA-DNA hybridization, protein profiles, fatty acid analysis, and nutritional profiles: A. indigens (type species) and A. communis (Reinhold-Hurek et al., 1993), A. tolulyticus (Zhou et al., 1995), A. toluclasticus and A. toluvorans (Song et al., 1999), A. evansii (Anders et al., 1995), A. anaerobius (Springer et al., 1998) and A. buckelii (Mechichi et al., 2002). One isolate (BH72) is different from these at the species level according to DNADNA similarity studies (≤25% DNA binding); however, a species name was not given because it is only a singular strain (Reinhold-Hurek et al., 1993). Numerous additional soil-borne strains are localized in the Azoarcus clade according to 16S rDNA sequence analysis (examples given in Fig. 1). Most have been isolated under conditions of denitrification with aromatic hydrocarbons as a C-source, but the lack of nutritional/physiological data and DNA-DNA similarity values does not yet allow a species assignment. The phylogenetic distances within the three clades in the Azoarcus/Thauera group are similar (Reinhold-Hurek and Hurek, 2000), and the soilborne and plant-associated species show considerable phenotypic differences (Reinhold-Hurek

Azoarcus spp. occur in terrestrial ecosystems; however, both groups of Azoarcus spp. strongly differ in their habitats and ecology. Members of A. indigens, A. communis and Azoarcus sp. BH72 occur inside roots or on the root surface of Gramineae and have never been isolated from root-free soil so far (Reinhold-Hurek and Hurek, 1998b), except for members of A. communis originating not from plants but from French refinery oily sludge (S2; Laguerre et al., 1987; Reinhold-Hurek et al., 1993) or from a compost biofilter in Canada (Lu1; Juteau et al., 1999). In contrast, all isolates localized in the clade of A. tolulyticus, A. toluvorans, A. toluclasticus, A. evansii, A. buckelii or A. anaerobius do not originate from living plants but mostly from soil and sediments. Therefore, their habitats will be treated separately. The plant-associated strains were originally isolated from Kallar grass (Leptochloa fusca [L.] Kunth), a flood-tolerant salt marsh grass grown as a pioneer plant on salt-affected, flooded, lowfertility soils in the Punjab of Pakistan since the 1970s (Reinhold et al., 1986; Reinhold-Hurek et al., 1993). These nitrogen-fixing bacteria were found in high numbers in surface-sterilized roots or culms and rarely on the root surface (A. communis). Only much later, the host range was found to be extended to rice (Oryza sativa) from Nepal (Engelhard et al., 2000) or to resting stages (sclerotia) of a plant-associated basidiomycete found in rice field soil from Pakistan (Hurek et al., 1997b). Soil-borne species of Azoarcus are very widespread but not plant-associated. Strains belonging to the validly described species have been isolated from noncontaminated soils (Song et al., 1999) or contaminated soils, either with unknown contaminations in industrial areas

876

B. Rareinhold-Hurek and T. Hurek

(Song et al., 1999) or with petroleum contaminations (Fries et al., 1994; Zhou et al., 1995). Many strains were also cultured from sediments of creeks (uncontaminated or contaminated; Anders et al., 1995), aquifers (Fries et al., 1994; Fries et al., 1997; Zhou et al., 1995), or even from activated sewage sludge (Springer et al., 1998). Soil-borne Azoarcus spp. are also widespread with respect to their geographical distribution: they have been found for example in North America (United States, Canada and Puerto Rico), South America (Brazil), or Europe (Germany and Switzerland).

Isolation Plant-associated and soil-borne members of Azoarcus spp. will be treated separately. Plantassociated strains are best enriched on media free of combined nitrogen. Washed or surfacesterilized roots are macerated aseptically in wash medium (enrichment medium free of a carbon source). To avoid overgrowth by faster growing diazotrophs, inoculate enrichment cultures with serial dilutions of this material. N-free semisolid synthetic malate medium (SM-N) is used for enrichment, composed of (per liter of distilled water): MgSO4 · 2H2O, 0.2 g; NaCl, 0.1 g; CaCl2, 0.02 g; MnSO4 · H2O, 0.01 g; Na2MoO4 · 2H2O, 0.002 g; Fe(III)-EDTA, 0.066 g; KH2PO4, 0.6 g, and K2HPO4, 0.4 g, adjusted to pH 6.8; DL-malic acid, 5 g, and KOH, 4.5 g adjusted to pH 6.8; agar, 2 g; and vitamin solution, 1 ml. The vitamin solution contains (mg per liter of distilled water) myo-inositol, 10,000; niacinamide, 100; pyridoxine HCl, 100; thiamine HCl, 100; calcium pantothenate, 50; folic acid, 20; choline chloride, 50; riboflavin, 10; ascorbic acid, 100; p-aminobenzoic acid, 1; vitamin A, 0.5; vitamin D3, 0.5; vitamin B12, 0.5; and D-biotin, 0.5. Add phosphate buffer and vitamin solution after autoclaving. Inoculate medium solidified in screw-cap tubes with 10 µl of macerate or root pieces below the medium surface. Incubate tubes without shaking at 30°C or 37°C, and check for development of a subsurface pellicle. Take samples for streaking before the pellicle has moved to the medium surface and become very dense. Isolate single colonies on SM-medium agar supplemented with 20 mg of yeast extract per liter. Use acetylene reduction method to analyze enrichment cultures or isolates for nitrogenase activity in semisolid medium. Check purity and colony color on complex medium, VM ethanol (which is based on SM medium but supplemented [per liter] with “Lab Lemco” powder [Oxoid] or Bacto Peptone [Difco], 3 g; yeast extract, 1 g; NaCl, 1 g; NH4Cl, 0.5 g; and agar, 15 g). Alternatively, for saltaffected habitats, use SSM medium which is

CHAPTER 3.2.18

adapted to high salt concentrations of salinesodic soil (Reinhold et al., 1986) instead of SM medium for enrichment and isolation. For soil-borne strains, enrichment is routinely done under strictly anaerobic conditions, with nitrate as terminal electron acceptor. A variety of aromatic carbon sources have been used, depending on the species or strain under study. Preparation of media and cultivation of bacteria have to be carried out under strictly anoxic conditions. Use a typical medium (for A. evansii), which consists of (per liter of distilled water): KH2PO4, 0.816 g; K2HPO4, 5.920 g; NH4Cl, 0.530 g; MgSO4 · 7H2O, 0.200 g; KNO3, 0.5 g; and CaCl2 · 2H2O, 0.025 g; dissolve phosphate and other ingredients separately from each other, adjust both to pH 7.8 for A. evansii or neutral pH for other strains, and combine both autoclaved solutions after cooling. Add 10 ml of sterile trace elements SL-10 and 5 ml of vitamin solution. SL-1 contains (per liter of distilled water): HCl (25%; 7.7 M), 10 ml; FeCl2·4H2O, 1.5 g; ZnCl2, 70 mg; MnCl2 · 4H2O, 100 mg; H3BO3, 6 mg; CoCl2 · 6H2O, 190 mg; CuCl2 · 2H2O, 2 mg; NiCl2 · 6H2O, 24 mg; and Na2MoO4 · 2H2O, 36 mg. First dissolve FeCl2 in HCl, then dilute in water, add and dissolve the other salts. The vitamin solution contains (mg per liter of distilled water): vitamin B12, 50 mg; pantothenic acid, 50 mg; riboflavin, 50 mg; pyridoxamine-HCl, 10 mg; biotin, 20 mg; folic acid, 20 mg; nicotinic acid, 25 mg; nicotinamide, 25 mg; α-lipoic acid, 50 mg; p-aminobenzoic acid, 50 mg; and thiamine-HCl · 2H2O, 50 mg. Add potentially toxic carbon sources at low concentrations (toluene, 5 ppm; phenol, 1 mM; and Nabenzoate, 5 mM) and spike cultures again with the carbon source after its degradation. Use ascorbic acid (4 mM) to reduce the medium. Carry out enrichment in anoxically sealed serum bottles. When using rich sediments as an inoculum, deplete when necessary from readily oxidizable carbon sources by repeated incubation in medium free of carbon sources (Fries et al., 1994). Plate the enriched samples on agar media such as M-R2A medium (Fries et al., 1994) or on agar used for dilution series (Widdel and Bak, 1992). M-R2A medium contains (per liter of distilled water): yeast extract, 0.5 g; peptone, 0.5 g; casamino acids, 0.5 g; glucose, 0.5 g; soluble starch, 0.5 g; sodium pyruvate, 0.3 g; K2HPO4, 0.4 g; KH2PO4, 0.25 g; KNO3, 0.505 g; CaCl2 · 2H2O, 0.015 g; MgCl2 · 2H2O, 0.02 g; FeSO4 · 7H2O, 0.007 g; Na2SO4, 0.005 g; NH4Cl, 0.8 g; MnCl2 · 4H2O, 5 mg; H3BO3, 0.5 mg; ZnCl2, 0.5 mg; CoCl2 · 6H2O, 0.5 mg; NiSO4 · 6H2O, 0.5 mg; CuCl2 · 2H2O, 0.3 mg; NaMoO4 · 2H2O, 0.01 mg; and agar, 15 g. Adjust the pH to 7.0 before autoclaving.

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

For cultivation on poorly water-soluble compounds such as alkylbenzenes, special procedures are required (Rabus and Widdel, 1995).

Identification Since members of the genus Azoarcus are quite heterogeneous with respect to ecology and physiology, the description of the genus is complex. The phenotypic characteristics of the genus Azoarcus are as follows: Cells are straight to slightly curved rods. Cell pairs often appear slightly S-shaped, with the exception of A. buckelii cells, which occur as cocci. Cell width is 0.4– 1.5 µm and length 1.1–4.0 µm. In many species, some elongated cells (8–12 µm) occur in late-log or stationary-phase cultures on medium containing malic acid and molecular nitrogen (N2) or ammonia as nitrogen source. Cells accumulate polyhydroxybutyrate granules and are Gram negative. Cells are highly motile by means of one, rarely two polar flagella. Many species fix nitrogen and then require microaerobic conditions for growth on N2. On semisolid nitrogenfree media, microaerophilic growth can be observed as veil-like pellicles developing several mm below the surface and moving to the medium surface during growth. Colony morphology differs in plant-associated and soilborne species. Azoarcus communis, A. indigens and Azoarcus sp. BH72 grow well on complex media such as trypticase soy agar (TSA) or VM medium containing beef extract or peptone, yeast extract, and ethanol (colony diameter 2– 4 mm after 4 d); on the latter medium, they produce a yellowish, nondiffusible pigment. In contrast, the soil-borne species A. tolulyticus, A. evansii, A. buckelii, A. toluvorans and A. toluclasticus grow poorly or not at all on complex media, and some may show different pigmentation. Azoarcus anaerobius is an exception since it is strictly anaerobic and therefore does not develop colonies on aerobically incubated agar A

Fig. 2. Microcopic images of Azoarcus spp. Transmission electron (A) and phase contrast (B, C) images. A) Azoarcus sp. strain BH72 cultured in liquid VM medium for 24 h. B) Azoarcus indigens VB32T cultured on N2 in semisolid SM medium for 24 h. C) Azoarcus tolulyticus BL2 grown on nutrient plates for 48 h. Bars, 5 µm. A) and B) are from Reinhold-Hurek et al. (1993) and C) is from Song et al. (1999).

C

877

plates. Optimum temperature for growth is 28–40°C, and no growth occurs at 45°C (Fig. 2). The soil-borne species are chemoorganoheterotrophic. Bacteria have a strictly respiratory metabolism with O2 as the terminal electron acceptor, except one species (A. anaerobius which is strictly anaerobic). Alternatively, under anaerobic conditions, nitrate can be used for dissimilatory nitrate reduction. Nitrate can be also assimilated. Bacteria are oxidase positive. They grow well on salts of organic acids (such as Lmalate, succinate, fumarate and DL-lactate), ethanol and L-glutamate, but not on mono- or disaccharides except for species that are not plant-associated. These soil-borne species use few carbohydrates and a variety of aromatic substrates as sole carbon sources under denitrifying conditions. Growth factor requirement varies: some strains depend on p-aminobenzoic acid or on cobalamin. All investigated species have 16:1, and all except one cis-9 16:1 and 18:1 as major cellular fatty acids. The mol% G+C of the DNA is 62–68 (Tm). The type species is Azoarcus indigens. Since the genera Azovibrio, Azospira and Azonexus were formerly included in Azoarcus sensu lato (Reinhold-Hurek and Hurek, 2000) and are physiologically very similar, some characteristics distinguishing these genera from each other are given in Table 1. Genera harboring plant-associated diazotrophs and Thauera were also included in Table 1. The genus Thauera (Macy et al., 1993; Anders et al., 1995), which is phylogenetically closely related, is physiologically similar to soil-borne species and thus difficult to distinguish by classical tests. The validly described species of Azoarcus are A. indigens (type species) and A. communis (Reinhold-Hurek et al., 1993), A. tolulyticus (Zhou et al., 1995), A. toluclasticus and A. toluvorans (Song et al., 1999), A. evansii (Anders et al., 1995), A. anaerobius (Springer et al., 1998) and A. buckelii (Mechichi et al., 2002). These species can be distinguished from each other B

β + 0.6–0.8 Beige − − − − − − − − − + − −

β da 0.4–1.0b Yellowish to beigec − da −

+f +g +h +i +k d d −l d

Subclass Cells curved Cell width (µm) Colony color

− − − − + + + − −

β + 0.4–0.6 Pinkish translucent − − −

Azospira

− − − − + − − − +

− − +

β + 0.6–0.8 Ocher

Azonexus

d + − − + − + + +

− + −

β + 0.6–0.7 Cream

Herbaspi-rillum

− + + + d + + + +

− + −

β − 0.3–0.8 Cream

Burkholderia vietnamiensis

ND ND − ND d ND + d +

α + 0.8–1.4 Pinkish opaque d + −

Azospirillum

a

Symbols and abbreviations: +, positive for all strains; −, negative for all strains; d, 11–89% of strains are positive; and ND, not determined. Positive for soil-borne species, negative for plant-associated species. b Except for A. anaerobius and A. buckelii (1.5 µm). c Several strains do not grow on nutrient agar. d Tested for T. aromatica. e Except for T. selenatis. f ND for A. toluvorans, A. toluclasticus, A. anaerobius, A. buckelii. g Negative for A. evansii. h Negative for A. toluclasticus, d for A. tolulyticus. i Negative for A. indigens, ND for A. toluvorans, A. toluclasticus, A. anaerobius. j ND for T. mechernichensis. k Negative for A. tolulyticus, ND for A. toluvorans, A. toluclasticus, A. anaerobius, A. buckelii. l ND for A. buckelii.

Fermentative Growth on sugars Requirement for cobalamine Growth in presence of O2 on n-Butylamine 4-Hydroxybenzoate Phenylacetate Glutarate 2-Oxoglutarate n-Caproate Proprionate D-Mannose L-Proline

Azovibrio

Azoarcus

Character

ND ND ND ND ND ND ND ND +

+ + −

α − 0.7–0.9 Brown

Gluconacetobacter diazotrophicus

Thauera

ND + + +j ND d +d ND +d

−d −e −

β − 0.7–1.0 White-yellowishc

Table 1. Differential characteristics of the genera Azoarcus, Azovibrio, Azospira, Azonexus and physiologically similar diazotrophs or related bacteria of the Proteobacteria.

878 B. Rareinhold-Hurek and T. Hurek CHAPTER 3.2.18

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

by phenotypic characteristics given in Table 2. Strain BH72 is included in Table 2 and may represent a species of its own according to DNADNA-similarity studies (Reinhold-Hurek et al., 1993). It has to be noted, however, that in some cases characteristics have been tested for one strain only (see also Table 3). A more complete list of the characteristics of these species is given in Table 3. Further comments on the species are given in the text below. Azoarcus indigens is the type species and can be differentiated from the others by its requirement for p-aminobenzoic acid, growth on itaconate, and a combination of characteristics given in Table 2. Cells are thin (0.5–0.7 µm wide) and curved, and cell pairs look slightly S-shaped. Colonies are very compact and difficult to disperse; growth in liquid media is clumpy; aggregation is very strong on peptone media. Optimum temperature for growth is 40°C. The major fatty acids are 3-OH-10:0, 12:0, cis-9 16:1, cyclo-17:0, and 18:1. Azoarcus indigens was isolated from roots and stem bases of Leptochloa fusca (L.) Kunth from the Punjab of Pakistan (ReinholdHurek et al., 1993), and black sclerotia of an Ustilago-related basidiomycete from rice soil in the Punjab of Pakistan (Hurek et al., 1997b) and from rice roots (Oryza sativa) from Nepal (Engelhard et al., 2000). The type strain is strain VB32 (=LMG 9092), which has a G+C content of 62.4 mol% (Tm). Accession number of the 16S rDNA sequence is AF011345 (www.ncbi.nlm. nih.gov/entrez/query.fcgi?db=nucleotide=search =AF011345). Azoarcus anaerobius can be distinguished from others by its strictly anaerobic life style with nitrate as the only electron acceptor. Nitrate is quantitatively reduced to N2 gas, nitrite accumulates intermediately, and nitrous oxide (N2O) is not detected. Additional differences are given in Table 2. Optimum growth temperature is 28°C. Enhanced salt concentration impairs growth. Cells are superoxidedismutase-positive. Sulfate, thiosulfate, sulfite, sulfur, trimethyl-amine N-oxide, dimethyl sulfoxide (DMSO), Fe(OH)3, K3[Fe(CN)6], or fumarate are not reduced, and oxygen is not reduced, not even at low pressures. Characteristics in addition to those given in Table 3 are: Sole carbon sources for growth are propanol, valerate, pyruvate, cyclohexane carboxylate, phenol, resorcinol, p-cresol; no growth with L-malate, formate, 5-oxocaproate, pimelate, catechol, hydroquinone, 2-hydroxybenzoate, ocresol, and m-cresol. No autotrophic growth with hydrogen or thiosulfate. The pH range is 6.5–8.2 and optimum pH 7.2. Cells were isolated from sewage sludge. The species consists of only one strain, which is the type strain LuFRes1 (= DSM 12081). It has a G+C content of 66.5 ±

879

0.5 mol% (Tm). Accession number of the 16S rDNA sequence is Y14701 (www.ncbi.nlm.nih. gov/entrez/query.fcgi?db=nucleotide=search= Y14701). Azoarcus buckelii can be distinguished from others by its cell shape and a combination of characteristics given in Table 2. Cells are coccoid of irregular size (1.5 ± 0.5 µm). Optimum temperature for growth is 28°C with a range from 4 to 40°C. It requires vitamin B12 for growth. Also nitrite can be used as terminal electron acceptor. Under oxic conditions, methanol and pyruvate are utilized in addition to carbon sources listed in Table 3. No growth occurs on formate, 2-aminobenzoate, catechol, pimelate, hexadecane, o-cresol, p-cresol, cyclohexane carboxylate, and phenol. Under denitrifying conditions, they utilize benzoate, 3-hydroxybenzoate, 4-hydroxybenzoate, 2aminobenzoate, o-phtalate, protocatechuate, phenol, o-cresol, p-cresol, cyclohexane carboxylate, 4-hydroxyphenylacetate, heptanoate, 2hydroxybenzaldehyde, 3-hydroxybenzaldehyde, 4-hydroxybenzaldehyde, 2-hydroxybenzoate, 3hydroxybenzoate, 4-hydroxybenzoate, 2,4dimethylphenol, 4-hydroxy-3-methylbenzoate, or for slow growth, 3-methylbenzoate, 2fluorobenzoate, gentisate, phenylacetate, phenylpropionate, L-tyrosine, L-phenylalanine, and indole 3-acetate; no growth occurs on m-cresol, 3-aminobenzoate, 4-aminobenzoate, 2methylbenzoate, 4-methylbenzoate, catechol, resorcinol, hydrochinone, cinnamate, p-anisate, L-tryptophan, indole, ethylbenzene, benzene, 3chlorobenzoate, adipate, pimelate, D-glucose, D-fructose, D-ribose, lactose, cyclohexanol, cyclohexanone, 1,3-cyclohexanedione, D,Lmandelate, D,L-4-hydroxymandelate, and 2hydroxybenzyl alcohol. The species consists of only one strain, which is the type strain U120 (= DSMZ 14744). It has a G+C content of 66 mol%. Accession number of the 16S rDNA sequence is AJ315676 (www.ncbi.nlm.nih.gov/ entrez/query.fcgi?db=nucleotide=search= AJ315676). Azoarcus communis can be distinguished from others by its cell width (0.8–1.0 µm) in combination with growth on D-mandelate, by growth on citrate, and a combination of characteristics given in Table 2. Cells are only slightly curved and plump. Optimum temperature for growth is 37°C. Some strains still grow well in 2% NaCl. The major fatty acids are 3-OH-10:0, 14:0, cis-9 16:1, 16:0, and 18:1. This species was isolated from roots of L. fusca (L.) Kunth from the Punjab of Pakistan (SWub3T; Reinhold-Hurek et al., 1993; Engelhard et al., 2000) and from refinery oily sludge in France (Laguerre et al., 1987). The species affiliation of strain LU1 isolated from a compost biofilter in Canada (Juteau et al., 1999)

0.8–1.0 + + + − + + + d + + − − − − − − − + + −

+ + + + − − − − − − − + + + − −

A. communis

0.5–0.7 +a + + +

A. indigens

d d d − d + + + + + + − − − − +

0.8–1 + d + −

0.6–0.8 + + + − + + − + + + − − − − − − − − − −

A. tolulyticus

A. sp. BH72

− + ± − − ± + + − − + ND ND ND ND +

0.8–1.0 + + ND −

A. toluvorans

− − d − − − + + d d + ND ND ND ND +

0.6–0.8 + − ND −

A. toluclasticus

+ d + + − + + d d + − − + − − −

0.4–0.8 + + − −

A. evansii

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

1.5 − − − −

A. anaerobius

Symbols and abbreviations: +, positive for all strains; −, negative for all strains; ±, weak reaction for all strains; d, 11–89% of strains are positive; and ND, not determined.

Cell width (µm) O2 terminal electron acceptor Catalase Nitrogen fixation Requirement for p-aminobenzoic acid Sole carbon sources for growth 3-Hydroxybenzoate Phenylacetate L-Phenylalanine D-Tartrate n-Caproate Isovalerate L-Proline, D(+)-galactose, sucrose Maltose D(+)-Glucose D(+)-Fructose D(−)-Ribose, adipate Itaconate p-Aminobenzoate D-Mandelate Citrate L(+)-Arabinose, D(+)-xylose

Characteristic

Table 2. Characteristics differentiating species of the genus Azoarcus from each other.

+ ± + ND − ND ND ND − − ND ND ND ND − ND

1.5 + + − −

A. buckelii

880 B. Rareinhold-Hurek and T. Hurek CHAPTER 3.2.18

Colony surface Growth at 40°C Requirement for p-aminobenzoic acid Requirement for cobalamin Nitrogen fixation Oxidation/fermentation of glucose Catalase Sole carbon sources for growthg n-Butylamine 3-Hydroxybenzoate Benzylamine Phenylacetate Benzoate L-Phenylalanine

Colony color (CR)c

Cell width (µm) Cell length (µm) Elongated cells in stationary cultures Colony diameter (mm, VM/CR)c Colony color (VM)c

Characteristic

2–4

Translucent yellowish Whiteish pink pink center Smooth + −

d

+ −

+

+ + v + d +

Opaque yellowish Whiteish pink white margin Rough + +

−

+ −

+

+ + d + d +

0.8–1.0 1.5–3.0 r

Azoarcus communis (n = 3)

2–3

0.5–0.7 2.0–4.0 r

Azoarcus indigens (n = 5)

+ + + + + −

+

+ −

−

Shining + −

Translucent yellowish Orange red

2–3/1

0.6–0.8 1.5–4.0 r

Azoarcus sp. BH72 (n = 1)

+b d +b d + d

D

+ ND

−

Shining −b −

Opaque yellowishb Orange redb

ND − ND + + ±

+

ND ND

−

ND − −

ND

ND

ND

0.8–1.0 1.4–2.8 ND

0.8–1b 1.4–2.8 − 1–1.5b

Azoarcus toluvorans (n = 2)

Azoarcus tolulyticusb (n = 9)

ND − ND − + d

−

ND ND

−

ND − −

ND

ND

ND

0.6–0.8 1.7–4.0 ND

Azoarcus toluclasticus (n = 5)

Table 3. Characteristics of species of the genera Azoarcus, Azovibrio, Azospira and Azonexus.a

+b +b +b d + +

+

+ +d

−

Shining −b −

Translucent beigeb Orange redb

0.2–0.7b

0.4–0.8 1.5–3.0 −

Azoarcus evansiib (n = 2)

ND + ND ± + +

+

− ND

+

ND + −

ND

ND

Shining d −

− − −

ND + ND + + +

−

− −

−

− − − − − −

+

+ −

−

Opaque beige Orange red

− −

1.5–2/1

0.6–0.8 1.5–3.5 r

Azovibrio restrictus (n = 5)

0

1.5 2.7–3.3 ND

1.5 ± 0.5 1.5 ± 0.5 − ND

Azoarcus anaerobius (n = 1)

Azoarcus buckelii (n = 1)

− − − − − −

ND

+ −

+

ND ND −

Negligible

Negligible

Negligible

0.6–0.8 1.5–3.5 ND

Azovibrio sp. BS20-3 (n = 1)

− − − − − −

+

+ −

d

Shining + −

Translucent pink-salmon Translucent orange

1–2

0.4–0.6 1.1–2.5 r

Azospira oryzae (n = 12)

− − − − − −

+

+ −

+

Shining + −

Opaque ocher Dark red

1–2/0.7

0.6–0.8 1.5–4.0 +

Azonexus fungiphilus (n = 5)

CHAPTER 3.2.18 The Genera Azoarcus, Azovibrio, Azospira and Azonexus 881

D(+) Malate L-Alanine 2-Oxoglutarate D-Tartrate n-Caproate Propionate Isovalerate L-Proline Malonate Maltose D(+)-Glucose D(+)-Fructose D(+)-Galactose, sucrose D(−)-Ribose Maltotriose, palatinose, D(+)-melezitose DL-Glycerate Toluene (denitrifying) Protocatechuate, L-mandelate 4-Hydroxybenzoate m-Coumarate Tryptamine Gentisate L(+)-Tartrate meso-Tartrate, betaineb p-Aminobenzoate Itaconate Glutarate D-Mandelate

Characteristic

Table 3. Continued

− − − − +

− −

− − +

+ + v + − − − − + +

− −

d − +

+ d + + − − + + − +

+f −b −b −b − − −b −b +b −b

+b + −b

+ +

+b +b −b − d d + +b − + + + +

Azoarcus tolulyticusb (n = 9)

ND d ND +f ND ND ND − ND ND ND ND ND

+f ND ND ND − ND ND ND ND ND

+ ND

ND ND ND − − − − + ND + d d +

Azoarcus toluclasticus (n = 5)

ND + ND

+ ND

ND ND ND − − − ± + ND + − − +

Azoarcus toluvorans (n = 2)

± ND ND − ND ND ND ND + ND

ND ND ND

+b ND −b − −b −b + +b +b + −b +b −b

ND ND

ND ND ND ND +f ND ND ND ND ND − − ND

Azoarcus buckelii (n = 1)

−b +b

−b +b +b + −b + +b + − d d +b +b

Azoarcus evansiib (n = 2)

+f ND ND ND ND ND ND ND ND ND

ND ND ND

ND ND

− ND ND ND ND + + ND ND ND − − ND

Azoarcus anaerobius (n = 1)

−b − − − − − − − − −

− − − − − − − − − −

− − −

− −

− − − − −

− + − − − + + − − − − − −

Azovibrio sp. BS20-3 (n = 1)

D D − − − + − − − − − − −

Azovibrio restrictus (n = 5)

− − − − − − − − − −

− − −

− −

− − + + + + + − − − − − −

Azospira oryzae (n = 12)

− − − − − − − − − −

− − −

− −

− − + − − − − + d − − − −

Azonexus fungiphilus (n = 5)

B. Rareinhold-Hurek and T. Hurek

+ + + + − − − − + −

+ + + + + + + − − − − − −

+ + + d + + + − − − − − −

+ d + + − + − − − − − − −

Azoarcus sp. BH72 (n = 1)

Azoarcus communis (n = 3)

Azoarcus indigens (n = 5)

882 CHAPTER 3.2.18

+ d d + v + + + − − − −

−

− − − − + + + + − − − −

−

−

− − − + + + + + − − − −

Azoarcus sp. BH72 (n = 1)

+

− − +b +b +b +b + + + − − +

Azoarcus tolulyticusb (n = 9)

d

ND ND ND ND + ND ND + + + − +

Azoarcus toluvorans (n = 2)

+

ND ND ND ND + ND ND d + + − +

Azoarcus toluclasticus (n = 5)

−

− − − +b + +b + + − − − −

Azoarcus evansiib (n = 2)

ND

− +h ND ND ± ND + ND ND ND ND ND

Azoarcus buckelii (n = 1)

ND

ND ND ND ND ND ND − + ND ND + −

Azoarcus anaerobius (n = 1)

−

− − − d + − + + − − − −

Azovibrio restrictus (n = 5)

−

− − − + − + + + − − − −

Azovibrio sp. BS20-3 (n = 1)

−

− − − v + + + + − − − −

Azospira oryzae (n = 12)

−

− − − − + + + + − − − −

Azonexus fungiphilus (n = 5)

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Symbols and abbreviations: +, positive for all strains; −, negative for all strains; ±, weak reaction for all strains; d, 11–89% of strains are positive; r, rarely; v, variable result; and ND, not determined. a All strains have the following features: cells are straight to curved rods except A. buckelii; oxidase positive; no growth in the presence of 5% NaCl and no growth-rate increase when NaCl is added to medium (ND for A. toluvorans, A. toluclasticus, A. anaerobius and A. buckelii); denitrification (ND for Azonexus sp.); no spore formation; no starch hydrolysis (ND for A. evansii, A. buckelii and A. anaerobius). b Marked characters were determined for A. tolulyticus strain Td-1 or A. evansii strain KB740T (Reinhold-Hurek and Hurek, 2000). c Growth on VM ethanol agar (VM) or Congo red agar (CR) at 37°C for 4 d. d Not tested for strain pF6. e All strains grow on DL-lactate, succinate, acetate, L-glutamate, butyrate (ND for A. buckelii), ethanol (ND for A. toluvorans and A. toluclasticus; no substrate but L-malate tested for A. evansii pF6); no growth on (ND for A. buckelii) D(+)-mannose, maltitol, N-acetyl-D-glucosamine, D-gluconate, and caproate; no growth onb (ND for A. toluclasticus, A. toluvorans, A. anaerobius and A. buckelii) D(+)-trehalose, L(+)-sorbose, ∀-D(+)melibiose, D(+)-raffinose, lactulose, 1-0-methyl-β-galactopyranoside, 1-0-methyl-∀-galactopyranoside, D(+) cellobiose, β-gentiobiose, 1-0-methyl-β-D-glucopyranoside, ∀-L-rhamnose, ∀-L-fucose, D(+)-arabitol, L(−)-arabitol, xylitol, dulcitol, D-tagatose, myo-inositol, D-sorbitol, adonitol, hydroxyquinoline-β-glucuronide, D-lyxose, erythritol, 1-0-methyl-∀-D-glucopyranoside, 3-0-methyl-glucopyranose, D-saccharate, mucate, D-glucuronate, Dgalacturonate, 2-keto-D-gluconate, 5-keto-D-gluconate, 3-phenylpropionate, caprylate, 4-aminobutyrate, DL-∀-amino-n-valerate, trigonelline, putrescine, ethanolamine, and D-glucosamine. f Under denitrifying conditions. g Tested with O2 as terminal electron acceptor. h Only glycerol tested. From Reinhold-Hurek et al. (1993), Anders et al. (1995), Hurek and Reinhold-Hurek (1995a), Zhou et al. (1995), Hurek et al. (1997), Rhee et al. (1997), Springer et al. (1998), Song et al. (1999), Engelhard et al. (2000), Reinhold-Hurek and Hurek (2000), and Mechichi et al. (2002).

Citrate Glycerol, D-alanine (−)-Quinate Isobutyrate L-Aspartate 3-Hydroxybutyrate L-Malate Fumarate Adipate L-Arginine L-Tyrosine L(+) Arabinose, D(+)-xylose ∀-Lactose

Characteristic

Azoarcus communis (n = 3)

Azoarcus indigens (n = 5)

CHAPTER 3.2.18 883

884

B. Rareinhold-Hurek and T. Hurek

is currently not clear. The type strain is strain SWub3 (= LMG 9095), which has a G+C content of 62.4 mol% (Tm). Accession number of the 16S rDNA sequence is AF011343 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=AF011343). Azoarcus evansii can be distinguished from other species by a combination of characters given in Table 2. Cells are rods with rounded ends and motile by means of a subpolarly inserted flagellum. Yeast extract (0.1% [w/v]) inhibits growth. Optimum growth temperature is 35–37°C, and optimum pH is 7.8. During denitrifying growth with aromatic compounds, nitrite is an intermediate and is reduced mainly to N2O (strain KB740T). Characteristics in addition to those listed in Table 3 are: Under anaerobic conditions, strain KB740T uses benzoate, phenylacetate, phenylglyoxylate, 3- and 4hydroxybenzoate, 2-aminobenzoate, 4-hydroxyphenylacetate, L-phenylalanine, p-cresol, 2fluorobenzoate, benzaldehyde, benzyl alcohol, indolylacetate, o-phtalate, adipate, pimelate, cyclohexane carboxylate, succinate, fumarate, Lmalate, acetate, acetone, D-fructose, D-maltose, and slow growth occurs on glutarate, D-glucose, but not on toluene, phenol, 2-hydroxybenzoate, protocatechuate, o- and m-cresol, indole, ethanol, D-ribose, or D-lactose. Pyridine is used by strain pF6 under aerobic and anaerobic conditions. The major cellular fatty acids are cis-9 16:1, 16:0, 12:0, 3-OH 10:0, and cis-9, 11 18:1. The major quinone is ubiquinone 8. The species description is based on one strain isolated from creek sediment in the United States (Braun and Gibson, 1984); however, a second strain, pF6, sharing 100% 16S rDNA sequence identity was isolated on pyridine from industrial wastewater in Korea (Rhee et al., 1997). The type strain is strain KB740 (= DSM6898), which has a G+C content of 67.5 mol%. Accession number of the 16S rDNA sequence is X77679 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=X77679). Azoarcus toluclasticus can be differentiated from the other species by the lack of catalase activity despite aerobic growth and by a combination of characters given in Table 2. Cells are short motile rods. Optimum temperature for growth is 30°C. Under denitrifying conditions, they can grow on acetate, benzoate, pyruvate, succinate, D-xylose, L-arabinose, D-ribose, Dgalactose, sucrose, lactose, maltose, adipate, lactate, mannitol, L-aspartate, L-proline or Larginine. All strains except strain MF23 can use toluene as a growth substrate and strains MF58 and MF63T can also grow on phenol under denitrifying conditions. The strains grow on brainheart infusion, nutrient and trypticase soy agar (TSA), except for strain MF63T, which does not

CHAPTER 3.2.18

grow on nutrient agar. The predominant cellular fatty acids are 16:0 and 16:1ω7c. DNA-DNA hybridizations show intermediate similarities (47–55%) among genomovar I (strains MF7 and MF23) and genomovar II (strains MF58, MF63T and MF441). They were isolated from aquifer sediments in the United States. The type strain is strain MF63 (= ATCC 700605), which has a G+C content of 67.3 mol% (HPLC). Accession number of the 16S rDNA sequence is AF123077 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=AF123077). Azoarcus tolulyticus can be differentiated from the other species by a combination of characters given in Table 2. Cells are short motile rods, which are slightly elongated (to 2.8 µm) when grown on M-R2A agar. Optimum temperature for growth is 30°C, and growth still occurs at 37°C. Under denitrifying conditions they can utilize acetate, adipate, L-arginine, L-arabinose, L-aspartate, benzoate, fumarate, D-galactose, Dglucose, lactate, lactose, maltose, mannitol, Lproline, pyruvate, D-ribose, succinate, sucrose, toluene, D-xylose, or 4-hydroxybenzoate. The strains grow on brain-heart infusion agar, but not or only poorly on nutrient and TSA. The predominant cellular fatty acids are 16:0 and 16: 1ω7c. Strains Td-19 and Td-3 (Zhou et al., 1995) have been removed from the species owing to low DNA-DNA similarity values (Song et al., 1999). The source of strains is aquifer sediments and petroleum-contaminated soils in the United States (Fries et al., 1994; Fries et al., 1997; Chee-Sanford et al., 1996). The type strain is strain Tol-4 (= ATCC 51758), which has a G+C content of 66.9 mol% (HPLC). Accession number of the 16S rDNA sequence is L33694 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=L33694). Azoarcus toluvorans can be differentiated from the other species by a combination of characters given in Table 2. Optimum temperature for growth is 30°C. Grow on brain-heart infusion, nutrient and TSA. Under denitrifying conditions, they can grow on acetate, benzoate, butyrate, fumarate, phenylacetate, pyruvate, succinate, toluene, D-xylose, L-arabinose, D-ribose, Dgalactose, sucrose, maltose, adipate, lactate, mannitol, L-aspartate, L-proline, L-phenylalanine, L-arginine, 4-hydroxybenzoate or phenol, and under aerobic conditions, on benzene or ethylbenzene. They were isolated from soil from an industrial area in Brazil (Td17) or noncontaminated organic soil in the United States (Td21T; Fries et al., 1994). The type strain is strain Td21 (= ATCC 700604), which has a G+C content of 67.8 mol% (HPLC). Accession number of the 16S rDNA sequence is L33692 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=L33692).

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Azoarcus sp. strain BH72 can be differentiated from the other species by its lack of growth on L-phenylalanine and a combination of characteristics given in Table 2. Cells are long, thin (0.6– 0.8 µm wide) and slightly curved, elongated (8–12 µm) cells occurring in late-log or stationary phase culture on N-free or ammoniumsupplemented SM-medium. Optimum temperature for growth is 40°C. The major fatty acids are16:0, cis-9 16:1, 18:1, 18:1, and 14:0. It was isolated from the root interior of Leptochloa fusca (L.) Kunth from the Punjab of Pakistan (Reinhold et al., 1986). The G+C content is 67.6 mol% (Tm). Accession number of the 16S rDNA sequence is AF011344 (www.ncbi.nlm. nih.gov/entrez/query.fcgi?db=nucleotide=search =AF011344).

Preservation Although organic acids are typical carbon sources for many strains, medium containing organic acids becomes alkaline in the stationary phase. Thus, subculturing on organic acids should be avoided and VM ethanol medium should be used instead. Soil-borne strains may also be subcultured on M-R2A medium. Preservation by lyophilization is possible. Long-term preservation is achieved by storage in liquid nitrogen with 5% DMSO as a cryoprotectant.

Physiology The optimum growth temperature is 30°C or 37– 40°C, which is lower for A. anaerobius (28°C). Optimum pH is neutral, and the bacteria are not halophilic. All Azoarcus species have a strictly respiratory type of metabolism, and no fermentative abilities have been observed. All species except one, A. anaerobius, use oxygen as the terminal electron acceptor. In the original description of Azoarcus spp., standard procedures had failed to reveal the capacity for denitrification (Reinhold-Hurek et al., 1993). In a closer examination, the capacity to use nitrate as the terminal electron acceptor was demonstrated for these plant-associated strains (Hurek and ReinholdHurek, 1995a), a feature which they share with all other species (Anders et al., 1995; Zhou et al., 1995; Springer et al., 1998; Song et al., 1999). Most strains belonging to A. tolulyticus, A. evansii, A. toluclasticus, A. toluvorans and A. anaerobius were enriched or isolated anaerobically on nitrate, whereas the plant-associated species were enriched under conditions of nitrogen fixation. In the original description of the genus Azoarcus, the capacity to fix nitrogen is listed as a genus character (Reinhold-Hurek et al., 1993). Nitrogen fixation (Fries et al., 1994; Zhou et al.,

885

1995) or the occurrence of a nitrogenase gene nifH (Hurek et al., 1997a) has also been demonstrated for A. tolulyticus. However, owing to the addition of new species, this cannot be generalized for Azoarcus anymore. Nitrogen fixation was not detected in A. anaerobius (Springer et al., 1998) or A. buckelii (Mechichi et al., 2002). In the absence of physiological data (Anders et al., 1995), A. evansii was thought unlikely to be diazotrophic since no polymerase chain reaction (PCR)-amplifiable nifH fragment (T. Hurek and B. Reinhold-Hurek, unpublished observations) or hybridization with a nifH probe (Hurek et al., 1997a) occurs. However, recently this species was reported to fix nitrogen (Mechichi et al., 2002). For most other isolates enriched under denitrifying conditions, this character was not tested at all. In all diazotrophic strains (A. indigens, A. communis, A. tolulyticus and Azoarcus sp. strain BH72), nitrogenase activity occurs only under microaerobic conditions, probably due to the lack of efficient oxygen protection mechanisms. It was shown that nitrogen fixation is more tolerant to oxygen in strain BH72 than in Azospirillum spp., reaching steady states in an oxygen-controlled chemostat until dissolved O2 exceeds 25 µM (Hurek et al., 1987). Similarly, the expression of nitrogenase genes is transcriptionally regulated in response to oxygen (fully repressed at 4% oxygen in the headspace) or combined nitrogen (repressed by 0.5 mM ammonium or nitrate; Egener et al., 1999). This strain shows augmented rates and efficiency of nitrogen fixation, called “hyperinduction,” in empirically optimized batch cultures when shifting down to extremely low oxygen concentrations (30 nM; Hurek et al., 1994a). In the course of hyperinduction, novel intracytoplasmic membrane stacks are formed which might participate in efficient nitrogenase activity, since the iron protein of nitrogenase is mainly associated with these membranes (Hurek and Reinhold-Hurek, 1995a). The formation of these so-called “diazosomes” is most abundant and reproducible in coculture of strain BH72 and endophytic Acremonium alternatum, a mitosporic deuteromycete related to the Hypocreales within the euascomycetes and an isolate from Kallar grass (Hurek et al., 1995b; Hurek and Reinhold-Hurek, 1998). As in many Proteobacteria, nitrogenase genes are organized in an nifHDK operon (Egener et al., 2001). Phylogenetically, the nitrogenase in the genus Azoarcus follows either the organismic phylogenetic tree or appears to have been obtained by lateral gene transfer, depending on the species. In A. indigens, A. communis and Azoarcus sp. strain BH72, a fragment of the iron protein of nitrogenase encoded by the nifH gene is most closely related to nitrogenases occurring in diazotrophs of the γ-subclass of Proteobacte-

886

B. Rareinhold-Hurek and T. Hurek

ria, while in A. tolulyticus, it is located within a clade of Alphaproteobacterial nitrogenases (Bradyrhizobium and Azospirillum; Hurek et al., 1997a). In general, carbohydrates are not the preferred carbon sources of Azoarcus spp. None of the plant-associated species is able to utilize any out of 50 mono- and disaccharides or sugar alcohols tested (Reinhold-Hurek et al., 1993). Azozrcus anaerobius shows no growth on common carbohydrates such as D(+)-glucose or D(–)fructose (Springer et al., 1998). In contrast, among the soil-borne species, all strains tested so far are able to utilize at least some carbohydrates (Table 1). All strains grow well on organic acids and a few amino acids (Table 1). The carbon sources listed here (except for A. anaerobius) were tested with O2 as the terminal electron acceptor. For some strains, tests were also carried out under anaerobic conditions with nitrate as electron acceptor. For the majority of carbon sources, results were identical; however, for several carbon sources, strain-dependent differences were found (Song et al., 1999). In contrast to plant-associated species (Reinhold-Hurek et al., 1993; Hurek and Reinhold-Hurek, 1995a), most soil-borne species grow on aromatic compounds (such as toluene or phenol) under denitrifying conditions (Zhou et al., 1995; Song et al., 1999), benzoate (Anders et al., 1995), or resorcinol (Springer et al., 1998). Owing to the anaerobic degradation of aromatic compounds, this bacterial group receives particular attention for their biodegradation and biotransformation abilities. While aerobic metabolism is characterized by the extensive use of molecular oxygen (which is essential for the hydroxylation and cleavage of the ring structures), different strategies are necessary for the anaerobic degradation and are currently under study. Toluene, which can be decomposed anaerobically by three species, is activated by the addition to fumarate to form benzylsuccinate (Beller and Spormann, 1997) catalyzed by benzylsuccinate synthase via a glycyl radical (Krieger et al., 2001). The aerobic metabolism of benzoate is unusual in A. evansii KB740T. None of the known pathways, including the conversion of benzoate to catechol (1,2-dihydroxybenzoene) or protocatechuate (3,4-dihydroxybenzoate), appears to operate in this species (Mohamed et al., 2001). The first step is the activation of benzoate to benzoyl-CoA by a benzoate-CoA ligase, and the second step involves the hydroxylation of benzoyl-CoA by a novel benzoyl-CoA oxygenase (Mohamed et al., 2001). The first step of phenylacetate degradation is catalyzed by two different phenylacetate-CoA ligases under aerobic and anaerobic conditions, respectively (Mohamed, 2000). Strain PF6 degrades pyridine

CHAPTER 3.2.18

aerobically and anaerobically with nitrate as electron acceptor (Rhee et al., 1997). BenzoylCoA is also a central intermediate in the anaerobic degradation of aromatic compounds (Harwood et al., 1998). In A. evansii KB740, nucleotide sequence analysis of the gene cluster including a gene for benzoate-CoA ligase indicates that the degradation of benzoate is probably similar to the benzoate-CoA pathway in Thauera aromatica (Harwood et al., 1998). For further reading on anaerobic oxidation of aromatic compounds, the review of Boll et al. (2002) is recommended. Resistance of Azoarus spp. to antibiotics has not been extensively tested. Azoarcus indigens, A. communis and strain BH72 are not resistant to ampicillin, kanamycin, streptomycin, spectinomycin or tetracycline (B. Reinhold-Hurek and T. Hurek, unpublished observation).

Genetics For the three plant-associated species of Azoarcus, DNA reassociation experiments led to the estimation of a genome size from 4.5 to 5 Mb. The genome of Azoarcus sp. strain BH72 is currently sequenced (Hurek and Reinhold-Hurek, 2003), with an estimated genome size of 4.4 Mb. Nothing is known about bacteriophages of the genus Azoarcus. Also the plasmid content is not well studied. In strain BH72, plasmids could not be detected in Eckhardt gels or by pulse field gel electrophoresis (B. Reinhold-Hurek and T. Hurek, unpublished observations). Several strains of Azoarcus spp. have been shown to be transformable. Broad host range plasmids based on RK2 such as pRK290 or pAFR3 can be transferred by triparental mating and are stably replicated in strain BH72 (Egener et al., 2001). Transformation can also be achieved by electroporation (Hurek et al., 1995b), and mutagenesis by allelic exchange (Hurek et al., 1995b) or transposon insertion (Dörr et al., 1998) is established.

Ecology As already stated in the section on Habitat, the two phylogenetic clusters of Azoarcus strains appear to differ substantially in their ecology: the plant-associated and the soil-borne species. Among the plant-associated species (A. indigens, A. communis and strain BH72), most strains (except strain S2 from sludge [Laguerre et al., 1987] or strain LU1 from a compost biofilter [Juteau et al., 1999], both related to A. communis [Fig. 1]) appear to be strictly plant-associated and have never been isolated from root-free soil (Reinhold-Hurek and Hurek, 1998b). In contrast, no members of soil-borne species have been isolated from plant material as yet.

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Plant-associated Azoarcus spp. have been up to now isolated from a limited number of samples, such as Kallar grass (Leptochloa fusca L. Kunth) from the Punjab of Pakistan (Reinhold et al., 1986; Reinhold-Hurek et al., 1993), rice roots (Oryza spp.) from Nepal (Engelhard et al., 2000), or resting stages of plant-associated fungi (Hurek et al., 1997b). However they may be more widely distributed than assumed; in molecular-ecological studies on root material or fungal spores, Azoarcus 16S rDNA genes (Hurek and Reinhold-Hurek, 1995a) or nifH genes (Ueda et al., 1995; Engelhard et al., 2000; Hurek et al., 2002) have been retrieved which did not correspond to genes of cultivated strains or species. Interestingly, the sequences retrieved up to now from plant material did not cluster with genes from soil-borne species, confirming that the latter do not appear to be plant associated. Since bacteria corresponding to the detected sequences could not be isolated from the same plant samples, they may occur in a state in which they are difficult to cultivate and thus overlooked by classical microbiological techniques. This is in agreement with the observation that in an inoculation experiment of Kallar grass with strain BH72, already two months after inoculation, attempts to reisolate the bacteria failed; however, nitrogenase gene (nifH) mRNA of this species was present at high levels in plant roots, suggesting high metabolic activity (Hurek et al., 2002). Using nif – and wild type BH72, it was demonstrated by N-balance and 15N2/14N2 isotope studies that Azoarcus sp. can supply fixed nitrogen to its host plant Kallar grass (Hurek et al., 2002), an important observation with respect to the ecological role and possible applications of these bacteria. Since plant-associated Azoarcus spp. cannot be isolated from root-free soil, the question arises how do they newly infect plants in nature. Survival in the absence of plants may be achieved in association with fungal resting stages in the soil (Hurek). Black sclerotia of a basidiomycete (a relative of Ustilago hordei) collected from a Kallar grass site in Pakistan harbored culturable strains of Azoarcus indigens and of the related genera Azovibrio spp., Azospira oryzae and Azonexus fungiphilus (Hurek). This suggests that plant-associated fungi might serve as shuttle vectors for plant-associated bacterial strains. Indeed, when these sclerotia were used to inoculate surface-sterilized rice seedlings in gnotobiotic culture, diazotrophs could be isolated from roots after 2–3 weeks of incubation. The abundant diazotrophs were identified as Azospira oryzae (B. Reinhold-Hurek, unpublished results). Mechanisms of plant-microbe-interactions are best studied for Azoarcus sp. strain BH72, orig-

887

inating from the pioneer plant Kallar grass from Pakistan (Reinhold et al., 1986). In gnotobiotic cultures in the laboratory, these bacteria have a wider host range. They can invade rice roots and to a lesser extent stems (Hurek et al., 1994b), where they mainly colonize the root cortex tissue intercellularly and rarely the stele including xylem cells (Hurek et al., 1994b). Despite a high density of colonization of the root interior, the bacteria do not cause symptoms of plant disease (Hurek et al., 1994b) and thus have an endophytic and not a pathogenic lifestyle. Unlike rhizobia, they do not form an endosymbiosis in living plant cells (Reinhold-Hurek and Hurek, 1998a). Nevertheless they show endophytic nitrogen fixation, expressing nitrogenase genes in the apoplast of aerenchymatic air spaces of flooded rice seedlings (Egener et al., 1999) or field-grown Kallar grass plants (Hurek et al., 1997a). Soil-borne species occur in a wide range of environments as stated in the section on Habitat. In most cases they were readily isolated under denitrifying conditions from nonmarine habitats such as contaminated or noncontaminated soils or sediments. Since many strains can degrade a wide range of aromatic compounds—which are the second most abundant class of organic compounds in nature—they can be expected to play an important role in carbon degradation in the biosphere. Their occurrence in anoxic sediments or sewage sludge indicates that their lifestyle in situ might be anaerobic (on nitrate as terminal electron acceptor), in contrast to the plantassociated strains that might be fixing nitrogen microaerobically. Besides the numerous isolates, there are also strains which have as yet escaped cultivation. One example is a deeply-branching clade of Azoarcus sp. found in high numbers in nitrifying-denitrifying activated sludge of an industrial wastewater treatment plant (Juretschko et al., 2002).

The Genera Azovibrio, Azospira and Azonexus Introduction Bacteria of these genera were first described to belong to the genus Azoarcus sensu lato (Reinhold-Hurek et al., 1993; Hurek et al., 1997b), to group C (Azovibrio), group D (Azospira) and group E (Azonexus). The availability of new isolates recently allowed a reassessment of the genus by a polyphasic taxonomic approach, leading to the proposal of three new genera. Since they are relatively similar in physiology and ecology, they are treated together.

888

B. Rareinhold-Hurek and T. Hurek

Recently, perchlorate-reducing bacteria were described as a new genus (Dechlorosoma suillum; Achenbach et al., 2001); however, a polyphasic taxonomic analysis by us (Tan and Reinhold-Hurek, 2003) demonstrated that they are members of Azospira oryzae. Therefore, these bacteria are also included in this chapter.

Phylogeny Analysis of almost complete 16S rDNA sequences placed all three genera into the Betaproteobacteria. Phylogenetic analyses using different algorithms showed that Azoarcus sensu lato was not monophyletic, but Azoarcus sensu stricto was divided from the other groups by the Thauera clade (Reinhold-Hurek and Hurek, 2000; see Fig. 1). The three new genera clearly represent different lineages, despite their exact phylogenetic relationships within the Rhodocyclus subgroup not being well resolved. Their phylogenetic distances from each other are similar to that of the Thauera/Azoarcus group or Duganella (formerly Zoogloea; Fig. 1). Other related genera are Dechloromonas, Propionivibrio and Rhodocyclus, and none are typically plantassociated bacteria.

Taxonomy The three groups are divided from each other and to Azoarcus by a polyphasic taxonomic analysis including 16S rDNA sequence analysis, DNA-DNA hybridization, protein profiles, fatty acid analysis, and nutritional profiles. Each genus consists of only one validly described species: Azovibrio restrictus, Azospira oryzae and Azonexus fungiphilus (Reinhold-Hurek and Hurek, 2000). The members of each species are quite homogeneous in protein patterns and substrate utilization, except for Azovibrio sp. strain BS203. It shares only 97% 16S rDNA sequence identity with Azovibrio restrictus S5b2T, which is strictly microaerobic and might thus deserve the rank of a separate species in future. Azovibrio now harbors new members recently described as a misnamed genus “Dechlorosoma suillum” (Achenbach et al., 2001). The type strain shares almost 100% of 16S rDNA sequence identity (1433 out of 1435 bp) with Azospira oryzae 6a3, is very similar in sodium dodecylsulfate (SDS)-soluble protein patterns and carbon source profiles, and shows approx. 90% binding in DNA-DNA-hybridization studies; thus the type strain of Dechlorosoma suillum and Azospira oryzae have to be regarded as the same species (Tan and Reinhold-Hurek, 2003).

CHAPTER 3.2.18

Habitat and Ecology Bacteria of all three genera occur in terrestrial habitats. Azovibrio and Azospira strains have typically been isolated from surface-sterilized plant roots and are thus considered to be diazotrophic endophytes of grasses. Members of both genera were found in roots of Kallar grass (Leptochloa fusca (L.) Kunth) in the Punjab of Pakistan (Reinhold-Hurek et al., 1993) and in roots of wild rice and cultured rice Oryza sativa in Nepal (Engelhard et al., 2000). Azospira oryzae also was found in roots of cultured rice in Italy (Engelhard et al., 2000) and wild rice in the Philippines (Hurek et al., 2000; Reinhold-Hurek and Hurek, 2000). Some strains of both genera and Azonexus fungiphilus were additionally found in another microniche, the resting stages (sclerotia) of a basidiomycete related to Ustilago hordei (Hurek et al., 1997b); they were collected in rice field soil at the site in the Punjab of Pakistan, where formerly Kallar grass had been grown. The finding that culturable bacteria appear to persist in fungal resting stages but not in soil suggests that plantassociated fungi may play a role as shuttle vectors for reinfection of plants by these diazotrophic endophytes. However, Azonexus fungiphilus was thought to be exclusively fungusassociated (Reinhold-Hurek and Hurek, 2000) but was recently cultured from surface-sterilized roots of wild rice, as well (B. Reinhold-Hurek and Z. Y. Tan, unpublished observations). A new group of Azospira oryzae (Tan and Reinhold-Hurek, 2003), which was misnamed “Dechlorosoma suillum” has an entirely different habitat. It was isolated from a primary treatment lagoon of swine waste in the United States (Achenbach et al., 2001).

Isolation Since all known strains are diazotrophic, enrichment and isolation are done on nitrogen-free media. The procedures are the same as described for plant-associated Azoarcus spp.

Identification Azovibrio, Azonexus and Azospira are quite similar in physiological features but differ especially in cell morphology. Characters differentiating them from Azoarcus and other related genera are given in Table 1. More detailed phenotypic features are listed in Table 3. All three genera are diazotrophic. They do not use carbohydrates as sole carbon sources, but they do use a few organic and amino acids. Additional features are given below.

CHAPTER 3.2.18 Fig. 3. Microscopic images of cells of the genera Azovibrio, Azospira and Azonexus. Transmission electron (A) and phase contrast (B-E) images of Azovibrio restrictus S5b2T (A, B), Azospira oryzae 6a3T (C), and Azonexus fungiphilus BS5-8T (D, E). Cells were grown on N2 in semisolid medium and photos taken during exponential (B), stationary (E) and late stationary (C, D) growth phase. Bars, 1 µm (A), 5 µm (B-D) or 10 µm (E). From Reinhold-Hurek and Hurek (2000).

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

A

889

B

C

D

Azovibrio cells are motile, slightly curved rods of 0.6–0.8 × 1.5–3.6 µm (Fig. 3), and elongated cells occur very rarely in stationary phase cultures. Cells grow well at 37°C. The major cellular fatty acids are cis-9-16:1, 16:0, 18:1, 14:0 and 3-OH-10:1. They have no dissimilatory nitrate reductase. The only validly described species is A. restrictus; however, strain BS20-3 (isolated from black sclerotia) might represent another species. In contrast to A. restrictus, BS20-3 requires cobalamin; it is also unusual because it is strictly microaerobic with negligible colony growth (Reinhold-Hurek and Hurek, 2000). The type strain is A. restrictus S5b2 (= LMG9099) which has a G+C content of 64.8 mol% and was isolated from surface-sterilized roots of Kallar grass. Accession number of the 16S rDNA sequence is AF011349 (www.ncbi.nlm.nih.gov/ entrez/query.fcgi?db=nucleotide=search= AF011349). Azospira cells are thin curved rods of 0.4–0.6 × 1.1–2.5 µm (Fig. 3), and elongated cells with one to several spiral windings of up to 8 µm occur rarely in late-stationary phase nitrogen-fixing cultures. Cells are highly motile with corkscrewlike motion. Growth is optimum at 37°C or 40°C. The major cellular fatty acids are cis-9-16:1, 16:0, 18:1 and 3-OH-10:1. They have no dissimilatory nitrate reductase. Plant-associated strains can use oxygen and nitrate as terminal electron acceptors, while strains originating from wastewater (formerly [Dechlorosoma suillum]) can also use perchlorate. The only validly described species is Azospira oryzae. The type strain is

E

strain 6a3 (= LMG9096) which has a G+C content of 65.2 mol% and was isolated from surfacesterilized roots of Kallar grass. Accession number of the 16S rDNA sequence is AF011347 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=AF011347). Azonexus cells are highly motile, slightly curved rods of 0.6–0.8 × 1.5–4.0 µm; elongated straight to coiled cells of up to 50 µm length occur in the stationary phase of nitrogen-fixing cultures (Fig. 3). Cells grow well at 37°C and require cobalamin for growth. The only validly described species is Azonexus fungiphilus. The type strain is strain BS5-8 (= LMG 19178), which was isolated from resting stages (black sclerotia) of a basidiomycete related to Ustilago found in a rice field in Pakistan. Accession number of the 16S rDNA sequence is AF011350 (www.ncbi. nlm.nih.gov/entrez/query.fcgi?db=nucleotide= search=AF011350).

Preservation Preservation and storage are carried out as for plant-associated species of Azoarcus.

Physiology All three genera (even the perchlorate-reducing isolates from swine wastewater) are diazotrophic, showing microaerophilic nitrogen fixation (Tan and Reinhold-Hurek, 2003). As the plant-associated Azoarcus strains, cells require microaerobic conditions for nitrogen fixation

890

B. Rareinhold-Hurek and T. Hurek

and form subsurface pellicles due to microaerophilic growth in semisolid media. The optimum growth temperature is 37–40°C. Optimum pH is neutral, and the bacteria are not halophilic. All genera have a strictly respiratory type of metabolism, and no fermentative abilities have been observed. All genera use oxygen or nitrate as the terminal electron acceptor, and some strains of Azospira oryzae also use perchlorate. All are oxidase and catalase positive. Similarly to plantassociated Azoarcus species, organic acids, amino acids, and ethanol but no carbohydrates are utilized.

Genetics Extrachromosomal genetic elements have not been intensively studied in these genera. Transposon mutagenesis has been established for Azonexus fungiphilus (Gemmer and B. Reinhold-Hurek, unpublished), while Azospira oryza appears to be difficult to transform.

Literature Cited Achenbach, L. A., U. Michaelidou, R. A. Bruce, J. Fryman, and J. D. Coates. 2001. Dechloromonas agitata gen. nov., sp. nov. and Dechlorosoma suillum gen. nov., sp. nov., two novel environmentally dominant (per)chloratereducing bacteria and their phylogenetic position. Int. J. Syst. Evol. Microbiol. 51:527–533. Anders, H. J., A. Kaetzke, P. Kämpfer, W. Ludwig, and G. Fuchs. 1995. Taxonomic position of aromatic-degrading denitrifying pseudomonad strains K 172 and KB 740 and their description as new members of the genera Thauera, as Thauera aromatica sp. nov., and Azoarcus, as Azoarcus evansii sp. nov., respectively, members of the beta subclass of the Proteobacteria. Int. J. Syst. Bacteriol. 45:327–333. Beller, H. R., and A. M. Spormann. 1997. Anaerobic activation of toluene and o-xylene by addition to fumarate in denitrifying strain T. J. Bacteriol. 179:670–676. Boll, M., G. Fuchs, and J. Heider. 2002. Anaerobic oxidation of aromatic compounds and hydrocarbons. Curr. Opin. Chem. Biol. 6:604–611. Braun, K., and D. T. Gibson. 1984. Anaerobic degradation of 2-aminobenzoate (anthranilic acid) by denitrifying bacteria. Appl. Environ. Microbiol. 48:102–107. Chee-Sanford, J. C., W. F. Frost, M. R. Fries, J. Zhou, and J. M. Tiedje. 1996. Evidence for acetyl coenzyme A and cinnamoyl coenzyme A in the anaerobic toluene mineralization pathway in Azoarcus tolulyticus Tol-4. Appl. Environ. Microbiol. 62:964–973. Dörr, J., T. Hurek, and B. Reinhold-Hurek. 1998. Type IV pili are involved in plant-microbe and fungus-microbe interactions. Molec. Microbiol. 30:7–17. Egener, T., T. Hurek, and B. Reinhold-Hurek. 1999. Endophytic expression of nif genes of Azoarcus sp. strain BH72 in rice roots. Molec. Plant-Microb. Interact. 12:813–819. Egener, T., D. E. Martin, A. Sarkar, and B. Reinhold-Hurek. 2001. Role of a ferrodoxine gene cotranscribed with the nifHDK operon in N2 fixation and nitrogenase “switch

CHAPTER 3.2.18 off” of Azoarcus sp. strain BH72. J. Bacteriol. 183:3752– 3760. Engelhard, M., T. Hurek, and B. Reinhold-Hurek. 2000. Preferential occurrence of diazotrophic endophytes, Azoarcus spp., in wild rice species and land races of Oryza sativa in comparison with modern races. Environ. Microbiol. 2:131–141. Fries, M., J. Zhou, J. Chee-Sanford, and J. M. Tiedje. 1994. Isolation, characterization, and distribution of denitrifying toluene degraders from a variety of habitats. Appl. Environ. Microbiol. 60:2802–2810. Fries, M. R., J. J. Forney, and J. M. Tiedje. 1997. Phenol- and toluene-degrading microbial populations from an aquifer in which successful trichloroethene cometabolism occurred. Appl. Environ. Microbiol. 63:1523–1530. Harwood, C., G. Burchhardt, H. Herrmann, and G. Fuchs. 1998. Anaerobic metabolism of aromatic compounds via the benzoyl-CoA pathway. FEMS Microbiol. Rev. 22:439–458. Hurek, T., B. Reinhold, I. Fendrik, and E. G. Niemann. 1987. Root-zone-specific oxygen tolerance of Azospirillum spp. and diazotrophic rods closely associated with Kallar grass. Appl. Environ. Microbiol. 53:163–169. Hurek, T., B. Reinhold-Hurek, G. L. Turner, and F. J. Bergersen. 1994a. Augmented rates of respiration and efficient nitrogen fixation at nanomolar concentrations of dissolved O2 in hyperinduced Azoarcus sp. strain BH72. J. Bacteriol. 176:4726–4733. Hurek, T., B. Reinhold-Hurek, M. van Montagu, and E. Kellenberger. 1994b. Root colonization and systemic spreading of Azoarcus sp. strain BH72 in grasses. J. Bacteriol. 176:1913–1923. Hurek, T., and B. Reinhold-Hurek. 1995a. Identification of grass-associated and toluene-degrading diazotrophs, Azoarcus spp., by analyses of partial 16S ribosomal DNA sequences. Appl. Environ. Microbiol. 61:2257– 2261. Hurek, T., M. Van Montagu, E. Kellenberger, and B. Reinhold-Hurek. 1995b. Induction of complex intracytoplasmic membranes related to nitrogen fixation in Azoarcus sp. BH72. Molec. Microbiol. 18:225–236. Hurek, T., T. Egener, and B. Reinhold-Hurek. 1997a. Divergence in nitrogenases of Azoarcus spp., Proteobacteria of the β-subclass. J. Bacteriol. 179:4172–4178. Hurek, T., B. Wagner, and B. Reinhold-Hurek. 1997b. Identification of N2-fixing plant- and fungus-associated Azoarcus species by PCR-based genomic fingerprints. Appl. Environ. Microbiol. 63:4331–4339. Hurek, T., and B. Reinhold-Hurek. 1998. Interactions of Azoarcus sp. with rhizosphere fungi. In: A. Varma and B. Hock (Eds.) Mycorrhiza. Springer-Verlag. Berlin, Germany. 595–614. Hurek, T., Z. Tan, N. Mathan, T. Egener, M. Engelhard, P. Gyaneshwar, J. K. Ladha, and B. Reinhold-Hurek. 2000. Novel nitrogen-fixing bacteria associated with the root interior of rice. In:J. K. Ladha and P. M. Reddy (Eds.) The Quest for Nitrogen Fixation in Rice. International Rice Research Institute. Los Banos, The Philippines. 47– 62. Hurek, T., L. Handley, B. Reinhold-Hurek, and Y. Piché. 2002. Azoarcus grass endophytes contribute fixed nitrogen to the plant in an unculturable state. Molec. PlantMicrob. Interact. 15:233–242. Hurek, T., and B. Reinhold-Hurek. 2003. Azoarcus sp. strain BH72 as a model for nitrogen-fixing grass endophytes. J. Biotechnol. 106:169–178.

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Juretschko, S., A. Loy, A. Lehner, and M. Wagner. 2002. The microbial community composition of a nitrifyingdenitrifying activated sludge from an industrial sewage treatment plant analyzed by the full-cycle rRNA approach. Syst. Appl. Microbiol. 25:84–99. Juteau, P., R. Larocque, D. Rho, and A. LeDuy. 1999. Analysis of the relative abundance of different types of bacteria capable of toluene degradation in a compost biofilter. Appl. Microbiol. Biotechnol. 52:863–868. Krieger, C. J., W. Roseboom, S. P. Albracht, and A. M. Spormann. 2001. A stable organic free radical in anaerobic benzylsuccinate synthase of Azoarcus sp. strain T. J. Biol. Chem. 276:12924–12927. Laguerre, G., B. Bossand, and R. Bardin. 1987. Free-living dinitrogen-fixing bacteria isolated from petroleum refinery oily sludge. Appl. Environ. Microbiol. 53:1674– 1678. Macy, J. M., S. Rech, G. Auling, M. Dorsch, E. Stackebrandt, and L. I. Sly. 1993. Thauera selenatis gen. nov. sp. nov., a member of the beta subclass of Proteobacteria with a novel type of anaerobic respiration. Int. J. Syst. Bacteriol. 43:135–142. Mechichi, T., E. Stackebrandt, N. Gad’on, and G. Fuchs. 2002. Phylogenetic and metabolic diversity of bacteria degrading aromatic compounds under denitrifying conditions, and description of Thauera phenylacetica sp. nov., Thauera aminoaromatica sp. nov., and Azoarcus buckelii sp. nov. Arch. Microbiol. 178:26–35. Mohamed, M. E.-S. 2000. Biochemical and molecular characterization of phenylacetate-coenzyme A ligase, an enzyme catalyzing the first step in aerobic metabolism of phenylacetic acid in Azoarcus evansii. J. Bacteriol. 182:286–294. Mohamed, M. E., A. Zaar, C. Ebenau-Jehle, and G. Fuchs. 2001. Reinvestigation of a new type of aerobic benzoate metabolism in the proteobacterium Azoarcus evansii. J. Bacteriol. 183:1899–1908. Rabus, R., and F. Widdel. 1995. Anaerobic degradation of ethylbenzene and other aromatic hydrocarbons by new denitrifying bacteria. Arch. Microbiol. 163:96–103. Reinhold, B., T. Hurek, E.-G. Niemann, and I. Fendrik. 1986. Close association of Azospirillum and diazotrophic rods with different root zones of Kallar grass. Appl. Environ. Microbiol. 52:520–526. Reinhold-Hurek, B., T. Hurek, M. Gillis, B. Hoste, M. Vancanneyt, K. Kersters, and J. De Ley. 1993. Azoarcus gen. nov., nitrogen-fixing proteobacteria associated with roots of Kallar grass (Leptochloa fusca (L.) Kunth) and

891

description of two species Azoarcus indigens sp. nov. and Azoarcus communis sp. nov. Int. J. Syst. Bacteriol. 43:574–584. Reinhold-Hurek, B., and T. Hurek. 1998a. Interactions of gramineous plants with Azoarcus spp. and other diazotrophs: Identification, localization and perspectives to study their function. Crit. Rev. Plant Sci. 17:29–54. Reinhold-Hurek, B., and T. Hurek. 1998b. Life in grasses: Diazotrophic endophytes. Trends Microbiol. 6:139–144. Reinhold-Hurek, B., and T. Hurek. 2000. Reassessment of the taxonomic structure of the diazotrophic genus Azoarcus sensu lato and description of three new genera and species, Azovibrio restrictus gen. nov., sp. nov., Azospira oryzae gen. nov., sp. nov, and Azonexus funguphilus gen. nov., sp. nov. Int. J. Syst. Evol. Microbiol. 50:649–659. Rhee, S. K., G. M. Lee, J. H. Yoon, Y. H. Park, H. S. Bae, and S. T. Lee. 1997. Anaerobic and aerobic degradation of pyridine by a newly isolated denitrifying bacterium. Appl. Environ. Microbiol. 63:2578–2585. Song, B., M. M. Haggblom, J. Zhou, J. M. Tiedje, and N. J. Palleroni. 1999. Taxonomic characterization of denitrifying bacteria that degrade aromatic compounds and description of Azoarcus toluvorans sp. nov. and Azoarcus toluclasticus sp. nov. Int. J. Syst. Bacteriol. 49:1129– 1140. Springer, N., W. Ludwig, B. Philipp, and B. Schink. 1998. Azoarcus anaerobius sp. nov., a resorcinol-degrading, strictly anaerobic, denitifying bacterium. Int. J. Syst. Bacteriol. 48:953–956. Tan, Z. Y., and B. Reinhold-Hurek. 2003. Dechlorosoma suillum Achenbach et al. 2001 is a later subjective synonym of Azospira oryzae Reinhold-Hurek and Hurek 2000. Int. J. Syst. Evol. Microbiol. 53:1139–1142. Ueda, T., Y. Suga, N. Yahiro, and T. Matsuguchi. 1995. Remarkable N2-fixing bacterial diversity detected in rice roots by molecular evolutionary analysis of nifH gene sequences. J. Bacteriol. 177:1414–1417. Widdel, F., and F. Bak. 1992. Gram-negative mesophilic sulfate-reducing bacteria. In: A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer. Berlin, Germany. 4:3352– 3378. Zhou, J., M. R. Fries, J. C. Chee-Sanford, and J. M. Tiedje. 1995. Phylogenetic analyses of a new group of denitrifiers capable of anaerobic growth on toluene and description of Azoarcus tolulyticus sp. nov. Int. J. Syst. Bacteriol. 45:500–506.

Index

A Absorption spectra Chromobacterium, 737 Rhodoferax, 596 Acetan production, Gluconacetobacter, 183–84 Acetate production Acetobacter, 176–77 Acetobacteraceae, 163 Acidomonas, 178 Asaia, 179–80 Gluconacetobacter, 180, 183 Kozakia, 190 phototrophic alphaproteobacteria, 55 Acetate utilization, phototrophic alphaproteobacteria, 56 Acetic acid bacteria. See Acetobacteraceae Acetic acid oxidation Acetobacter, 175–76 Acidomonas, 179 Gluconacetobacter, 181 Kozakia, 190 Acetobacter, 13, 151, 163–66, 168–70, 175–78. See also Gluconacetobacter applications of, 178 characteristics of, 168, 176–77 classification of, 167, 175–76 genetics of, 177–78 habitats of, 176 identification of, 176 isolation and cultivation of, 176 pathogenicity of, 178 taxonomy of, 175–76 Acetobacter aceti, 166, 168, 170–71, 174–78 Acetobacter acidophilus, 171 Acetobacter europaeus, 172 Acetobacter indonesiensis, 169 Acetobacter lovaniensis, 169, 173, 177 Acetobacter malorum, 177 Acetobacter oboediens, 172 Acetobacter orleanensis, 173 Acetobacter pasteurianus, 166, 168–73, 175–78, 731 Acetobacter peroxydans, 166, 176, 178 Acetobacter pomorum, 176–77 Acetobacter suboxydans, 365

Acetobacter tropicalis, 169 Acetobacter xylinum, 168, 171 Acetobacteraceae, 13, 163–90 Acetobacter, 13, 163–66, 175–78 Acidiphilium, 13 Acidomonas, 13, 163–66, 178–79 Asaia, 13, 163–66, 179–80 characteristics of, 168 classification of, 167 Gluconacetobacter, 13, 163–66, 180–85 Gluconobacter, 13, 163–66, 185–89 habitats and uses of, 163, 166–75 isolation, cultivation and preservation of, 175 Kozakia, 163–66, 189–90 morphology of, 163 phylogenetic tree of, 163–65 phylogeny of, 163–64 Rhodopila, 13 taxonomy of, 164–66 Acetone degradation, Xanthobacter, 308–9 Acetylmethylcarbinol production, Gluconobacter, 189 Achromobacter, 648, 678–89 applications of, 687–89 arsenite resistance by, 686 compound degradation by, 683–84 detection of, 686–87 heavy metal resistance by, 686 as human pathogens, 678–80 physiology of, 679 Achromobacter cycloclastes, 680–81, 687 Achromobacter denitrificans, 678 Achromobacter EST4002, 685 Achromobacter insolitus, 678 Achromobacter lyticus M497-1, 688 Achromobacter piechaudii, 678 Achromobacter ruhlandii, 678 Achromobacter sp. WM111, 686 Achromobacter spanius, 678 Achromobacter strain ABIV, 685 Achromobacter strain M-30-Y, 686 Achromobacter strain RS9, 685 Achromobacter xylosoxidans, 675, 678, 682–83, 686–87, 747 Achromobacter xylosoxidans subsp. denitrificans, 685 Achromobacter xylosoxidans subsp. xylosoxidans B3, 686 Achromopeptidase, 688

Acid pollution, Thiobacillus, 815 Acid production, Thiobacillus, 816, 819 Acidianus, 815 Acidiphilium, 13, 41, 163, 562, 565–66, 576, 580 Acidiphilium acidophilum, 819, 821 Acidiphilium rubrum, 571, 580 Acidisphaera rubrifaciens, 163 Acidithiobacillus, 18, 822–24 Acidithiobacillus albertis, 816, 822–23 Acidithiobacillus caldus, 823 Acidithiobacillus ferrooxidans, 815–16, 818–19, 822–24 Acidithiobacillus thiooxidans, 816, 822–24 Acidocella, 163 Acidomonas, 13, 163–66, 178–79 applications of, 179 characteristics of, 168, 179 classification of, 167 genetics of, 179 habitats of, 178 identification of, 178–79 isolation and cultivation of, 178 taxonomy of, 178 Acidomonas methanolica, 178–79 Acidophile Acidithiobacillus, 822–24 Beijerinckia, 151–52 Thiobacillus, 812 Acidovorax, 16, 723 Acidovorax facilis, 16 Acinetobacter, 20 Acremonium alternatum, 885 Actinobacillus, 19 Actinobacillus actinomycetemcomitans, 844–45 Actinomyces, 845 Activated sludge Achromobacter/Alcaligenes strains, 684–85 Alcaligenes, 677 Alcaligenes strains, 682 Azoarcus, 876 bulking of, 772 Comamonas, 726, 732 Leptothrix cholodnii, 768 Leptothrix lopholea, 768 Leptothrix-Sphaerotilus group, 763

894

Index

nitrite-oxidizing bacteria, 863, 865, 868 phototrophic betaproteobacteria, 596 Sphaerotilus, 772–73 Adenylate cyclase toxin, 662–64, 666, 668 Bordetella, 658, 660 Adherence Bordetella, 656–57, 667 Eikenella corrodens, 840 Adhesins Bordetella, 655–56, 665 Opa proteins as, 611 as toxins, 657 Aerobic phototrophic bacteria, 562–80 applications of, 580 ecology of, 580 genetics of, 578–80 habitats of, 565–66 identification of, 568–71 isolation of, 566–68 physiology of, 572–78 oxygen dependence and heterotrophic metabolism, 575–78 photosynthetic potential, 572–75 preservation of, 571 properties of, 563–64, 567 taxonomy, phylogeny and origin, 562–65 Aerocavin, 741 Aerocyanidin, 741 Aeromonadaceae, 19–20 Aeromonas, 19 Aeromonas, 19, 740 Aeromonas salmonicida, 19 Afipia, 15 African tick-bite fever, Rickettsia, 503 Agave habitat, Zymomonas, 201 Agracin sensitivity, Agrobacterium, 107, 109 Agrobacterium, 13–15, 91–110, 151, 315, 321, 324, 350, 365, 467–68, 659, 747, 749 applications of, 109–10 characteristics of, 94 disease from, 91–110, 105–9 binding to plant, 107–9 control of, 106–9 ecology of, 105 genetics of, 96–105 chromosomal, 97–101 general, 96–97 habitats of, 92 identification of, 92–94 isolation of, 92 morphology of, 94 phylogeny of, 91 physiology of, 94–96 preservation of, 94 taxonomy of, 91–92 Agrobacterium radiobacter, 91, 107, 109, 468 Agrobacterium rhizogenes, 91–92, 95, 101, 106, 109–10

Agrobacterium rubi, 91 Agrobacterium tumefaciens, 28, 91–92, 94–103, 105–10, 127–28, 130, 324, 371–72 Agrobacterium vitis, 91–92, 101, 104–6 Albamycin sensitivity, Methylobacterium, 262 Alcaligenaceae, 16–17, 675, 678, 689, 747 Alcaligenes, 16, 648, 675–77, 723, 754, 849 applications of, 687–89 arsenite resistance by, 686 compound degradation by, 682–86 denitrification by, 680–82 detection of, 686–87 heavy metal resistance by, 686 as human pathogens, 678–80 physiology of, 679 strains of, 681–88 Alcaligenes defragrans, 675, 677, 684 Alcaligenes denitrificans, 681, 683, 685–87 Alcaligenes denitrificans subsp. denitrificans, 689 Alcaligenes denitrificans subsp. xylosoxidans, 678, 682 Alcaligenes faecalis, 365, 675, 677, 680–83, 686–87, 689, 751, 868 Alcaligenes faecalis subsp. faecalis, 675, 677 Alcaligenes faecalis subsp. parafaecalis, 675, 677, 688 Alcaligenes latus, 675, 758–59 Alcaligenes xylosoxidans, 680–87, 850 Alcaligenes xylosoxidans subsp. denitrificans, 685, 688 Alcaligenes xylosoxidans subsp. xylosoxidans, 688 Alcalilimnicola, 19 Alcohol dehydrogenase isoenzymes, Zymomonas, 206–7 Alcohol utilization Acidomonas, 179 Agrobacterium, 91–92 Azospirillum, 115 Gluconobacter, 187 Herbaspirillum, 145–46 Kozakia, 190 Methylobacterium, 257 Paracoccus, 238, 240–43 phototrophic alphaproteobacteria, 56 Xanthobacter, 305–6 Algal habitat Aquaspirillum, 711 dimorphic prosthecate bacteria, 76–77, 86 Alkane degradation, Xanthobacter, 307–8 Alkanesulfonate degradation, Achromobacter/Alcaligenes strains, 684 Alkene degradation, Xanthobacter, 307–8 Allorhizobium, 14–15 Alphaproteobacteria, 3–29

Acetobacteraceae, 13 Anaplasmataceae, 13 Bartonellaceae, 14–15 Brucellaceae, 14–15 Caulobacteraceae, 14 ecology of, 9–15 Ehrlichiaceae, 13 group 1, 43–44 group 2, 44 group 3, 44 Hyphomicrobiaceae, 15, 290 morphology of, 9–15 orders and families of, 12 phototrophic, 41–59 phylogeny of, 9–15 physiology of, 9–15 Rhizobiaceae, 14–15 Rhizobiales, 294 Rhodospirillaceae, 13–14 Rickettsiaceae, 13 Rickettsiales, 457–64 Sphingomonadaceae, 14 Alteromonadaceae, 19–20 Alteromonas, 737 Alteromonas luteoviolacea, 740, 742 Alysiella, 18, 828–38 ecology of, 837–38 habitat of, 832–34 identification of, 835–37 isolation of, 834–35 morphology of, 828, 830, 834–36 phylogeny of, 828–32 preservation of, 837 taxonomy of, 832 traits of, 832 Alysiella filiformis, 828, 830–33, 836–37 Amaricoccus, 14 Amino acid synthesis Achromobacter/Alcaligenes strains, 687 Comamonas, 729 Thiobacillus, 819 Amino acid utilization aerobic phototrophic bacteria, 576 Brucella, 362, 366 dimorphic prosthecate bacteria, 82–83 epsilonproteobacteria, 24 Neisseria, 608 Paracoccus, 240–43 phototrophic alphaproteobacteria, 56 D-Aminoacylases, Achromobacter/Alcaligenes strains, 687 1-Aminocyclopropane-1-carboxylate deaminase, Alcaligenes strains, 682 Aminoglycoside sensitivity Brucella, 428–29 dimorphic prosthecate bacteria, 74 Ammonia assimilation, phototrophic betaproteobacteria, 598 Ammonia-oxidizing bacteria, 778–806, 862, 864, 866 characteristics of, 779–86

Index detection of, 794–800 classical techniques, 794–95 immunological techniques, 795 molecular techniques, 796–800 distribution in nature, 800–803 betaproteobacteria, 801–3 gammaproteobacteria, 803 enrichment, isolation, and maintenance of, 778–79 environmental populations, 803–6 genera of, 779–80 habitats of, 781 morphology of, 780, 783–86 phylogenetic tree of, 787–90 phylogeny of, 786–94 species of, 780–86 Ammonium utilization Azospirillum, 129 dimorphic prosthecate bacteria, 74 Ampicillin resistance, Xanthobacter, 303 Amylosucrase, Neisseria, 624 Anaerobiospirillum, 19–20 Anaplasma, 13, 320, 323, 460–61, 473, 493–94, 507–12, 547 ecology of, 510 epidemiology of, 510–11 genetics of, 509–10 identification of, 508–9 isolation of, 508 phylogeny of, 507–8 physiology of, 509 preservation of, 509 Anaplasma bovis, 493, 508 Anaplasma centrale, 511 Anaplasma marginale, 460–61, 493, 508–12 Anaplasma ovis, 508 Anaplasma phagocytophila, 493–94, 508–10 Anaplasma platys, 508–9, 511, 514 Anaplasmataceae, 13, 493 Anaplasma, 13, 493 Ehrlichia, 493 habitats of, 508 morphology of, 493 Neorickettsia, 493 phylogenetic tree of, 508 taxonomy of, 508 Wolbachia, 493, 547 Anaplasmosis Anaplasma, 508–12 control and prevention of, 511 vaccine for, 511–12 Ancalomicrobium, 65–70 characteristics of, 69 cultivation and maintenance of, 68–69 habitats of, 65–66 identification of, 69–70 isolation of, 66–68 morphology of, 65 phylogenetic tree of, 67 species of, 67 taxonomy of, 65 Ancalomicrobium adetum, 65–66, 68–69 Angiococcus, 22

Angiogenic factor, Bartonella, 472–73 AniA, 613 Neisseria, 607 Aniline degradation, Comamonas, 729 Animal disease Aeromonadaceae, 19 Anaplasma, 511 Anaplasmataceae, 493 Bartonella, 462, 467 Betaproteobacteria, 16 Brucella, 315, 335–38, 372–429 Campylobacteraceae, 24–25 Cardiobacteriaceae, 20 Enterobacteriaceae, 19 Francisella, 20 Gammaproteobacteria, 18 Neisseriaceae, 18 Pasteurellaceae, 19 Piscirickettsia, 20 Animal habitats Neisseria, 603 Simonsiella, 829 Antarctobacter, 14 Antibiotic production Chromobacterium, 741 Myxobacteria, 22 Antibiotic resistance/sensitivity Achromobacter/Alcaligenes strains, 680, 689 Azoarcus, 886 Bartonella, 484 Bordetella, 653 Brucella, 321, 350, 411–12, 428–29 Burkholderia, 852, 856 Chromobacterium, 739 dimorphic prosthecate bacteria, 74 Methylobacterium, 262 Neisseria, 624–25, 626, 630, 632 Rickettsia, 506 Spirillum, 707 Xanthobacter, 303 Antibiotic treatment gonorrhea, 632 meningococcal disease, 632 pertussis, 651, 653 Antibody response, to pertussis, 665–66 Antigenic structure, Eikenella corrodens, 842 Antigens Burkholderia, 854, 855 Neisseria, 617 Antimicrobial peptides, 664 Bordetella, 661–62 Antimicrobial susceptibility, Eikenella corrodens, 844 Antipyrin degradation, Phenylobacterium, 250–56 Appendicitis, Bilophila, 23 Appendix, Comamonas, 726 Apple habitat, Acetobacteraceae, 172–73 Applications Acetobacter, 178 Achromobacter/Alcaligenes strains, 687–89

895

Acidomonas, 179 aerobic phototrophic bacteria, 580 Agrobacterium, 109–10 Azospirillum, 132–34 Beijerinckia, 161 Brucella, 429–31 Chromobacterium, 740–41 Comamonas, 732 Coxiella, 541 Derxia, 756 dimorphic prosthecate bacteria, 85–86 Gluconacetobacter, 185 Gluconobacter, 188–89 Herbaspirillum, 147 Janthinobacterium, 743 manganese-oxidizing bacteria, 229 methanotrophs, 284 Methylobacterium, 263 Ochrobactrum, 749 Paracoccus, 246 phototrophic alphaproteobacteria, 58–59 phototrophic betaproteobacteria, 599 Phyllobacterium, 749 Proteobacteria, 12 Spirillum, 707–8 Wolbachia, 516, 555–56 Xanthobacter, 307–9 Aquabacter, 290, 294 Aquabacter spiritensis, 294 Aquabacterium, 758 Aquaspirillum, 701, 703, 710–21, 723 differentiating characteristics of, 712–13 enrichment of, 711–18 habitat of, 711 identification of, 719–21 isolation of, 711–19 maintenance of, 711–18 morphology of, 719 motility of, 719 preservation of, 718–19 rRNA superfamily III species of, 719–20 rRNA superfamily IV species of, 720–21 taxonomy of, 719–21 type strains of, 714 Aquaspirillum anulus, 720 Aquaspirillum aquaticum, 711, 717, 719, 720, 725–26 Aquaspirillum autotrophicum, 710, 711, 717, 720, 742–43 Aquaspirillum bengal, 710, 717, 719 Aquaspirillum delicatum, 710, 719, 720 Aquaspirillum dispar, 719, 720 Aquaspirillum fasciculus, 710, 716, 719, 720 Aquaspirillum giesbergeri, 720 Aquaspirillum gracile, 710, 716, 719, 720 Aquaspirillum itersonii, 141, 703, 710, 719, 720 Aquaspirillum itersonii subsp. itersonii, 720

896

Index

Aquaspirillum itersonii subsp. nipponicum, 718, 720 Aquaspirillum itersonii subsp. vulgatum, 720 Aquaspirillum magnetotacticum, 710, 717, 718, 719, 720 Aquaspirillum metamorphum, 715, 720 Aquaspirillum peregrinum, 719, 720 Aquaspirillum peregrinum subsp. integrum, 718 Aquaspirillum peregrinum subsp. peregrinum, 710 Aquaspirillum polymorphum, 710, 719, 720 Aquaspirillum psychrophilum, 710, 718, 720 Aquaspirillum putridiconchylium, 720 Aquaspirillum serpens, 710, 716, 717, 719, 720 Aquaspirillum sinuosum, 720 Aquaspirillum voronezhense, 701 Archangiaceae, 22 Archangium, 22 Arcobacter, 24–25 Arcobacter nitrofigilis, 25 Aromatic amine deamination, Alcaligenes strains, 682 Aromatic amine dehydrogenase, Alcaligenes strains, 682 Aromatic compound degradation Achromobacter/Alcaligenes strains, 683–84 Azoarcus, 873, 886 Comamonas, 728 phototrophic betaproteobacteria, 598 Arphamenines A and B, 741 Arsenite resistance, Achromobacter/Alcaligenes strains, 686 Arthrobacter globiformis, 731 Arthropod habitat Alcaligenes, 677 Anaplasma, 508–9 Ancalomicrobium, 66 Bartonella, 479 Coxiella, 530, 536 dimorphic prosthecate bacteria, 77 Ehrlichia, 512–13 Prosthecomicrobium, 66 Rickettsia, 496, 500 Wolbachia, 515–16, 547, 550–51, 553 Asaia, 13, 163–66, 172–73, 179–80 characteristics of, 168, 180 classification of, 167 habitats of, 180 identification of, 180 isolation and cultivation of, 180 taxonomy of, 180 Asaia bogorensis, 180, 190 Asaia krungthepensis, 180 Asaia siamensis, 180 Asticcacaulis, 72–86 Asticcacaulis biprosthecum, 74, 81, 84

Asticcacaulis excentricus, 74, 81, 84 ATP synthesis, Brucella, 367 Atrophic rhinitis, 651, 664 Autophagosomes, Brucella, 407–8 Autotransporter proteins, Rickettsia, 498–99 Auxotyping, Neisseria, 628 Azo dye degradation, Pigmentiphaga kullae, 689 Azoarcus, 17, 126, 873–88 characteristics of, 878 ecology of, 886–87 genetics of, 886 habitat of, 875 identification of, 877–85 isolation of, 876–77 phylogeny of, 873–75 physiology of, 885–86 preservation of, 885 species of, 880–83 taxonomy of, 875 Azoarcus anaerobius, 873, 875, 877, 879, 885–86 Azoarcus buckelii, 873, 875, 877, 879, 885 Azoarcus communis, 873, 875, 877, 879–86 Azoarcus evansii, 873, 875–77, 884–86 Azoarcus indigens, 873, 875, 877, 879, 885–87 Azoarcus sp. strain BH72, 873, 875, 877, 879, 885–87 Azoarcus toluclasticus, 873, 875, 877, 884–85 Azoarcus tolulyticus, 873, 875, 877, 884–86 Azoarcus toluvorans, 873, 875, 877, 884–85 Azoarcus/Thauera branch, 873–75, 888 Azomonas, 21, 154–55, 751, 753–54 Azomonas agilis, 160 Azonexus, 873, 877, 887–90 characteristics of, 878 genetics of, 890 identification of, 888–89 isolation of, 888 morphology of, 889 phylogeny of, 888 physiology of, 889–90 preservation of, 889 species of, 881–83 taxonomy of, 888 Azonexus fungiphilus, 887–90 Azorhizobium, 290, 294 Azorhizobium caulinodans, 294 Azospira, 17, 873, 877, 887–90 characteristics of, 878 genetics of, 890 identification of, 888–89 isolation of, 888 morphology of, 889 phylogeny of, 888 physiology of, 889–90 preservation of, 889 species of, 881–83 taxonomy of, 888 Azospira oryzae, 887–90

Azospirillum, 42, 115–34, 141, 151, 753–54 applications of, 132–34 characteristics of, 124 cultivation of, 122–23 habitats and ecology of, 116–20 history of, 115 identification of, 123–26 morphology of, 115, 121 phylogenetic tree of, 116 physiology of, 126–32 ammonium uptake and assimilation, 129–30 nitrogen fixation, 129 osmotolerance, 131 plant growth regulating substances, 130–31 plant root colonization, 127–28 siderophores, 131–32 preservation of, 123 species of, 118, 124 taxonomy of, 115–16 Azospirillum amazonense, 115–16, 121, 123, 125–26, 128–29, 131–32 Azospirillum brasilense, 115–16, 118–20, 123, 125, 127–33, 146–47, 720 Azospirillum doebereinerae, 115–16, 120, 124–25 Azospirillum halopraeferens, 115–16, 121–22, 125, 131 Azospirillum irakense, 115–16, 122–23, 126, 131–33 Azospirillum largimobile, 116, 122, 125, 132 Azospirillum lipoferum, 115–16, 118, 120, 122–25, 128–29, 131–32, 754 Azotobacter, 21, 154–56, 751, 753–54, 756 Azotobacter acida. See Beijerinckia indica Azotobacter chroococcum, 160, 275 Azotobacter vinelandii, 146, 160 Azotomonas insolita, 751 Azovibrio, 873, 877, 887–90 characteristics of, 878 genetics of, 890 identification of, 888–89 isolation of, 888 morphology of, 889 phylogeny of, 888 physiology of, 889–90 preservation of, 889 species of, 881–83 taxonomy of, 888 Azovibrio restrictus, 888, 889 Azurins, 681

B Bacillary angiomatosis, Bartonella, 480–81, 483–84 Bacillary peliosis, Bartonella, 481, 483–84 Bacillus extorquens, 257 Bacillus faecalis alcaligenes, 723

Index Bacillus subtilis, 861 Bacitracin resistance, Methylobacterium, 262 Bacteremia Bartonella, 462, 482–84 Coxiella, 536–37 Bacteriochlorophyll aerobic phototrophic bacteria, 562, 571, 575 Alphaproteobacteria, 9, 13 Bacteriochlorophyll a aerobic phototrophic bacteria, 562, 566 Methylobacterium, 263 phototrophic alphaproteobacteria, 49, 54 Bacteriochlorophyll b, phototrophic alphaproteobacteria, 50, 54 Bacteriochlorophyll β, phototrophic alphaproteobacteria, 50 Bacteriophage Acidomonas, 179 Brucella, 346–48, 359 Neisseria, 626 Bacteriophage production, Bartonella, 471–73 Bacteriovorax, 23 Bacteroides corrodens, 840 Bacteroides ureolyticus, 840 Balneatrix alpica, 21 Bartonella, 15, 315, 320, 323, 462, 467–85, 493 disease from, 480–84 antimicrobial therapy, 484 clinical presentation, 480–83 diagnosis, 483–84 epidemiology of, 479–80 genetics of, 478–79 habitats of, 469 identification of, 469–71 immunity from, 484–85 isolation of, 469 morphology of, 467, 470 pathogenesis host adherence, 474–75 host colonization, 473–74 host invasion, 475–78 phylogenetic tree of, 468 phylogeny of, 467 physiology of, 471–78 angiogenic factor, 472–73 cell structure, 471 enzymes, 473 extracellular products, 471–72 growth and metabolism, 471 pathogenesis, 473–78 preservation of, 471 taxonomy of, 467–69 Bartonella alsatica, 462 Bartonella bacilliformis, 15, 462, 467, 469, 471–75, 477–79, 482, 484 Bartonella clarridgeiae, 462, 470–71, 474, 480, 484 Bartonella doshiae, 462 Bartonella elizabethae, 462, 470–72, 475, 478–80, 484 Bartonella grahamhii, 462 Bartonella henselae, 15, 462, 467–76, 478–81, 483–85

Bartonella koehlerae, 462 Bartonella peromysci, 462 Bartonella quintana, 15, 462, 467–73, 475, 477–80, 482–85 Bartonella talpae, 462 Bartonella taylorii, 462 Bartonella tribocorum, 462, 478 Bartonella vinsonii, 471, 478 Bartonella vinsonii subsp. arupensis, 462 Bartonella vinsonii subsp. berkhoffii, 462 Bartonella vinsonii subsp. vinsonii, 462 Bartonella washoensis, 462 Bartonella weissii, 462 Bartonellaceae, 14–15, 467–68 Bartonella, 15, 467–68 Bartonellae, 462 Bartonellosis, Bartonella, 479–80, 484 Basidiomycete Azoarcus, 887 Azonexus, 888 Bdellovibrio, 23 Bdellovibrio group, 23 Bacteriovorax, 23 Bdellovibrio, 23 Bee hive environment Acetobacteraceae, 173–74 Gluconobacter, 186 Simonsiella, 832 Zymomonas, 201 Beer Acetobacter, 176 Acetobacteraceae, 172 Gluconobacter, 186 Zymomonas, 201 Beggiatoa, 20, 815 Beijerinckia, 15, 151–61, 751–56 applications of, 161 cultivation of, 154 habitats of, 151–52 identification of, 154–57 isolation of, 152–54 life cycle of, 155 morphology of, 151–52, 154–59 physiology of, 159–60 preservation of, 154 species differentiation, 157–59 Beijerinckia derxii, 154, 157, 159 Beijerinckia derxii subsp. derxii, 159 Beijerinckia derxii subsp. venezuelae, 159 Beijerinckia fluminensis, 154, 156–59 Beijerinckia indica, 152–54, 156–59 Beijerinckia indica subsp. indica, 158 Beijerinckia indica subsp. lacticogenes, 156, 158 Beijerinckia indica var. alba, 158 Beijerinckia mobilis, 154–58 Benzoate degradation, Azoarcus, 886 Betaproteobacteria, 3–29. See also Phototrophic betaproteobacteria Alcaligenaceae, 16–17 ammonia-oxidizing bacteria, 783–86

897

Comamonadaceae, 16–17 ecology of, 15–18 morphology of, 15–18 Neisseriaceae, 18 orders and families of, 16 phototrophic, 593–99 phylogeny of, 15–18 physiology of, 15–18 sulfur oxidizing bacteria of, 812–14, 821–22 Bilophila, 23 Bilophila wadsworthia, 23 Biochemistry Acetobacter, 176–77 Asaia, 180 Bartonella, 470 Cardiobacterium hominis, 844–45 Derxia, 755–56 Eikenella corrodens, 842 Gluconobacter, 186–87 Kozakia, 190 Neisseria, 606 Xanthobacter, 301–3 Biocontrol, Burkholderia, 848 Biodegradation Azoarcus, 886 Betaproteobacteria, 15–17 nitrite-oxidizing bacteria, 868–69 Biofilms Leptothrix-Sphaerotilus group, 763 nitrite-oxidizing bacteria, 863–65, 867 Biology, for speciation, 334–37 Bioremediation aerobic phototrophic bacteria, 580 Burkholderia, 848 Comamonas, 732 methanotrophs, 266, 284 Methylobacterium, 263 Pseudomonas, 21 Xanthobacter, 308–9 Biosynthesis, Neisseria, 620–21 Biphenyl degradation Achromobacter/Alcaligenes strains, 683 Comamonas, 730 Biphenyl/chlorobiphenyl dioxygenase, Comamonas, 730 Bird habitat Bordetella, 651 Coxiella, 530 Blastochloris, 44 Blastochloris sulfoviridis, 44, 46, 50, 54, 57 Blastochloris viridis, 44, 50, 56 Blood Achromobacter, 678 Alcaligenes, 677 Aquaspirillum, 711 Comamonas, 726 Ochrobactrum, 748 Bordetella, 16, 648–68, 675, 678 autotransporter protein family in, 656–57 description of, 648 diseases from, 650–51, 651 ecophysiology of, 653–62

898

Index

evolution of, 650 habitats of, 651–53 immunity to, 664–66 intracellular state of, 667–68 isolation of, 651–53 LPS of, 661–62 phylogeny of, 649–50 significance of, 650–51 species of, 648–49 toxins of, 658–61 vaccine resistance of, 666–67 virulence factors of, 656–58, 667–68 Bordetella avium, 648, 651, 655, 660 Bordetella bronchiseptica, 648–50, 653–55, 661–62, 668 Bordetella bronchiseptica cluster, 648, 652, 660–61, 663–64 characterization of, 649–50 virulence factors of, 667–68 Bordetella hinzii, 648 Bordetella holmesii, 648–49 Bordetella parapertussis, 648–51, 661–62, 667 Bordetella pertussis, 16, 351, 648–56, 659–60, 662–64, 666–68 toxins of, 658–61 transcriptional responses to, 664 virulence factors of, 656–58 Bordetella petrii, 648 Bordetella trematum, 648 Boutonneuse fever, Rickettsia, 503 Bovine habitat Anaplasma, 510–11 Anaplasmataceae, 493 Bartonella, 469 Brucella, 335, 372–82, 394, 429 Campylobacteraceae, 24–25 Coxiella, 530, 536 Succinivibrionaceae, 19–20 Brachymonas, 723 Brackiella oedipodis, 16 Bradyrhizobium, 13, 15, 320, 864 Bradyrhizobium japonicum, 754 Brenneria, 19 Brevundimonas, 14, 18 Brucella, 9, 13, 315–431, 467–68, 747, 845 applications of, 429–31 as adjuvants, 431 antitumoral and antiviral activity, 430 as biological weapon, 431 foreign antigen delivery, 430 as immunological tools, 431 characteristics of, 339 classification of, 322–23, 344–46 disease from, 381–429 control of infection, 411–15 diagnosis of, 415–28 pathobiology of, 387–411 pathology of, 381–87 treatment of, 428–29 epidemiology of, 372–81 control, eradication and prevention, 380 geographical distribution of, 373–75 host infection, 375

vaccines and vaccination, 375–80 zoonosis, 380–81 genetics of, 327–28, 368–72 chromosomes, 368–71 genome, 368 habitats of, 315, 335, 337, 349 identification of, 325–26, 341–48 brucellaphages, 346–48 cellular and colonial characteristics, 341–44 identification and typing, 344–46 isolation of, 337–41 phylogenetic tree of, 316–17, 321 phylogeny of, 315–30 ancestor-descendant relationships, 315–20 gene acquisition, 330 genome evolution, 321–24 speciation of, 320 virulence structure derivation, 324–29 physiology of, 349–68 metabolism, 361–68 structure, 349–61 preservation of, 348–49 taxonomy of, 330–37 genetic composition of, 332–33 species definition, 333–37 Brucella abortus, 320, 324, 329–31, 335–39, 341, 343, 345, 348–56, 359–68, 370–71, 374, 377–80, 382–85, 388, 390–97, 400–405, 409, 412–13, 417–18, 422, 424–25, 429–30 Brucella canis, 320, 330–31, 334–37, 339, 344–45, 353–54, 359, 365–66, 368, 374, 380, 382, 385, 390, 415, 418, 426–27 Brucella melitensis, 28, 320, 330–31, 335–39, 341, 343–45, 348, 352–54, 356, 359–60, 363–68, 372–74, 376–77, 379–80, 382–83, 385, 388, 408, 413, 418, 422, 425–26, 429–31 Brucella neotomae, 330–31, 335–37, 339, 345, 348, 368, 374, 380, 382, 385 Brucella ovis, 42, 330–31, 334–39, 341, 343, 345, 353–54, 365–66, 368, 371, 374, 379–80, 382, 385, 390, 415, 418, 426–27 Brucella suis, 320–21, 330–31, 335–39, 344–45, 348, 364–65, 368, 370, 372, 374, 378, 380, 382, 385, 388, 418, 431 Brucellaceae, 14–15, 840 Brucellosis Brucella, 372–75, 381–429 control of infection, 411–15 antibody response, 411–12 T cell response, 412–15 diagnosis of, 415–28 in animals, 416–17 antibody detecting tests, 421–24 cell-mediated immunity tests, 425–28 in humans, 415–16

immunological tests, 417–21 pathobiology of, 387–411 host invasion, 387 replication in host cell, 393–401 survival outside host cells, 387–93 virulence mechanisms, 401–11 pathology of, 381–87 in animals, 382, 385–86 in fetus, 382–85 in humans, 385 male infections, 385 outcome and self cure, 386–87 treatment of, 428–29 in animals, 429 in humans, 428–29 Buchnera, 19, 463 Buchnera aphidicola, 19, 28, 463 Budding dimorphic prosthecate bacteria, 73, 82 Methylobacterium, 261 Methylosinus, 280, 282 phototrophic alphaproteobacteria, 42–44 Seliberia, 585 Burkholderia, 16–18, 21, 848–56 disease of, 852–53 epidemiology of, 851–52 habitat of, 850 identification of, 850–51 isolation of, 850 pathogenicity of, 853–56 taxonomy of, 849–50 treatment of, 856 Burkholderia caryophilli, 849 Burkholderia cepacia, 16–17, 28, 848–54, 856 Burkholderia cepacia complex, 678, 849–52 Burkholderia gladioli, 849 Burkholderia mallei, 16, 848–49, 851–53, 855–56 Burkholderia multivorans, 850–52 Burkholderia picketii, 849 Burkholderia pseudomallei, 16, 848–56 Burkholderia solanaceareum, 849 Burkholderia stabilis, 849–52 Burkholderia thailandensis, 848–49 Burkholderia vietnamensis, 849–52

C Cadmium resistance Achromobacter/Alcaligenes strains, 686 Comamonas, 731 Caedibacter, 462 Caedibacter caryophila, 462 Calvin cycle, Paracoccus, 235 Campylobacter, 24–25, 703 Campylobacter jejuni, 24, 703 Campylobacteraceae, 24–25 Arcobacter, 24–25 Campylobacter, 24–25 Dehalospirillum, 24–25 Sulfurospirillum, 25

Index Canine habitat Anaplasma, 511 Anaplasmataceae, 494 Bartonella, 469, 479 Ehrlichia, 460 Neorickettsia, 461 Succinivibrionaceae, 19–20 Canine monocytotropic ehrlichiosis, Ehrlichia, 514–15 Capnogtophaga, 844 Caprine habitat Brucella, 372–82, 384, 394 Coxiella, 530, 536 Carbofurane degradation, Achromobacter/Alcaligenes strains, 686 Carbohydrate metabolism, Neisseria, 608 Carbohydrate transport, Zymomonas, 203–4 Carbon dioxide production, phototrophic alphaproteobacteria, 55 Carbon dioxide requirement Brucella, 366–67 Neisseria, 607–8 Carbon dioxide utilization ammonia-oxidizing bacteria, 778 phototrophic alphaproteobacteria, 56 Xanthobacter, 304–5 Cardiobacteriaceae, 20 Cardiobacterium, 20 Dichelobacter, 20 Suttonella, 20 Cardiobacterium, 20 Cardiobacterium hominis, 20, 844–45 Carnimonas, 20 Carotenoids aerobic phototrophic bacteria, 570–71, 575 Azospirillum, 125 phototrophic alphaproteobacteria, 44, 49 Rhodocyclus, 594–96 Caryophanon, 835 Cat flea typhus, Rickettsia, 503–4 Cat scratch disease, Bartonella, 462, 480, 483–84 Catalase production Brucella, 368 Coxiella, 541 dimorphic prosthecate bacteria, 74 Methylocystis, 280 Methylosinus, 280 Neisseria, 607 Catechol degradation, Comamonas, 728, 730 Catenococcus thiocyclus, 813, 817 Catheters, Comamonas, 726 Cats. See Feline habitat Caulobacter, 65, 72–86, 315 Caulobacter crescentus, 84, 370 Caulobacter leidyi, 77, 84 Caulobacter vibrioides, 83–84 Caulobacteraceae, 14 Brevundimonas, 14 CDC group Vd, 747, 748

Cell wall composition Bartonella, 471 Coxiella, 531 dimorphic prosthecate bacteria, 82 Methylococcus, 275 Orientia, 506 Rickettsia, 457 Xanthobacter, 302 Zymomonas, 203 Cell wall degradation, Achromobacter/Alcaligenes strains, 688 Cellulose production Agrobacterium, 97, 109 Gluconacetobacter, 183–85 Cellvibrio, 21 Cepabactin, Burkholderia, 853 Cepacia syndrome, 848 Cephalothin resistance, Methylobacterium, 262 Chemoprophylaxis, for meningococcal disease, 632 Chemostat culture, Thiobacillus, 817–19 Chemotaxis Agrobacterium, 96, 106 Azospirillum, 127 Xanthobacter, 301 Chicken habitat Alysiella, 832 Simonsiella, 832 Chlamydia, 463 Chlamydia pecorum, 463 Chlamydia pneumoniae, 463 Chlamydia psittaci, 463 Chlamydia trachomatis, 463 Chlamydiae, 463 Chlamydia, 463 morphology of, 463 Chloramphenicol resistance/ sensitivity Rickettsia, 506 Xanthobacter, 303 Chlorhexidine solutions, Achromobacter, 678 Chloridazon degradation, Phenylobacterium, 250–56 Chlorinated compound degradation, Achromobacter/Alcaligenes strains, 685 Chromate-reducing bacteria, Comamonas, 731 Chromatiaceae, 18–19 Chromatibacteria, 15 Chromatium, 703 Chromobacter violaceum, 851 Chromobacteriosis, 739 Chromobacterium, 18, 720, 737–41, 747 applications of, 740–41 differentiating characteristics of, 738 enrichment of, 739–40 habitat of, 738 identification of, 740 isolation of, 739–40 morphology of, 738

899

motility of, 738 preservation of, 740 Chromobacterium amethystinum, 741 Chromobacterium fluviatile, 737, 739–40 Chromobacterium lividum, 737 Chromobacterium membranaceum, 741 Chromobacterium violaceum, 737–42, 751 Chromohalobacter, 20 Chromosomes Agrobacterium, 91, 96–101, 110 Brucella, 321–24, 368–71 Coxiella, 535 Paracoccus, 245–46 Chronic granulomatous disease, 852 Cider Acetobacter, 176 Acetobacteraceae, 169–70 Gluconobacter, 186 Zymomonas, 201 Citrate utilization, phototrophic alphaproteobacteria, 56 Citric acid cycle, Phenylobacterium, 255 Citromicrobium, 562, 565, 571, 576 Citromicrobium bathyomarinum, 565–66, 572–73 Clarithromycin sensitivity, Orientia, 507 Clostridium, 688 Clostridium beijerinckii, 688 Cocoa wine Acetobacter, 176 Acetobacteraceae, 169 Gluconobacter, 186 Zymomonas, 201 Co-divergence hypothesis, 830 Coenzyme M, Xanthobacter, 308 Coffee plant habitat Acetobacteraceae, 173 Gluconacetobacter, 181, 184 Collectins, 663 Colonization Neisseria, 603, 605–6, 610 of oral cavity Alysiella, 833–34 Simonsiella, 833–34 Colony morphology Acidomonas, 179 Actinobacillus actinomycetemcomitans, 845 aerobic phototrophic bacteria, 571 Agrobacterium, 92 Alysiella, 835–37, 837 Azoarcus, 877 Azospirillum, 121–26 Bartonella, 469 Beijerinckia, 155, 157–59 Brucella, 341–44 Burkholderia, 850 Cardiobacterium hominis, 844 Derxia, 753–54, 755–56 Eikenella corrodens, 841–42 Haemophilus aphrophilus, 845 Kingella, 845

900

Index

Leptothrix, 768–70 Leptothrix-Sphaerotilus group, 762 Methylobacter, 278 Methylocaldum, 277 Methylococcus, 275 Methylocystis, 280 Methylomicrobium, 279 Methylomonas, 277 Methylosinus, 280 Phenylobacterium, 253 Simonsiella, 835–37, 837 Sphaerotilus, 773 Xanthobacter, 294–95, 300 Colorless sulfur bacteria. See Thiobacillus Comamonadaceae, 16–17, 723–24, 726, 758 Comamonas, 6, 16, 18, 720, 723–32 applications of, 732 characteristics of, 727 cultivation of, 728 ecology of, 731 genetics of, 729–31 habitat of, 726 history of, 725 identification of, 726–28 isolation of, 726 morphology of, 726 pathogenicity of, 731–32 phylogenetic tree of, 724 phylogeny of, 723 physiology of, 728–29 preservation of, 728 strain of, 728–32 taxonomy of, 723–26 Comamonas acidovorans, 723, 725–27, 850 Comamonas compransoris, 725 Comamonas denitrificans, 723, 726, 728 Comamonas nitrativorans, 723, 726, 728 Comamonas percolans, 723, 725 Comamonas terrigena, 723, 725–26, 728, 731–32 Comamonas testosteroni, 723, 725–32 Compost biofilter, Azoarcus, 875 Conferva ochracea, 758 Conjugation, Neisseria, 626 Contagious abortion, Brucella, 372 Contagious equine metritis, 690 Corroding colonies Eikenella corrodens, 840–45 species capable of, 843–45 Corrosion of agar, 843–45 Eikenella corrodens, 841–42 Thiobacillus, 815 Corynebacterium, 749 Cow manure, Aquaspirillum, 711 Cowdria, 13, 460 Cowdria ruminantium, 460 Cowdriosis. See Heartwater disease Coxiella, 21, 462, 529–41 applications of, 541 cultivation of, 533–34 epidemiology of, 536

genetics of, 535 habitats of, 530 identification of, 531–33 isolation of, 530–31 morphology of, 529 pathogenicity of, 536–41 phylogeny of, 529–30 physiology of, 534–35 Coxiella burnetii, 462, 493, 529–41 Craurococcus, 562, 566 Craurococcus roseus, 163, 562, 565 Crown gall tumors Agrobacterium, 91, 94, 105, 109 biocontrol of, 106–9 Cultivation Acetobacter, 176 Acetobacteraceae, 175 Acidithiobacillus ferrooxidans, 823–24 Acidomonas, 178 Ancalomicrobium, 68–69 Asaia, 180 Azospirillum, 122–23 Beijerinckia, 154 Brucella, 338–41 Comamonas, 728 Coxiella, 533–34 Derxia, 752–53 dimorphic prosthecate bacteria, 83–85 Gluconacetobacter, 181 Gluconobacter, 186 Kozakia, 190 manganese-oxidizing bacteria, 226–28 methanotrophs, 272–74 Methylobacterium, 259–61 Phenylobacterium, 251–52 Prosthecomicrobium, 68–69 Spirillum, 703–6 Thiobacillus, 816–19 Xanthobacter, 299–300 Zymomonas, 202 Cycloclasticus, 20 Cystic fibrosis, 852 Achromobacter, 678, 680 Burkholderia, 848, 849, 850, 851, 856 Cystobacter, 22 Cystobacteraceae, 22 Cysts Beijerinckia, 156–57 Methylobacter, 278, 281–82 Methylocaldum, 275–77 Methylocystis, 282 Methylomonas, 277 Cytochrome a, Beijerinckia, 160 Cytochrome c Alphaproteobacteria, 13 Beijerinckia, 160 Gluconacetobacter, 183 Gluconobacter, 186–87 Paracoccus, 243–44 Zymomonas, 211–12 Cytochrome c oxidase production Methylocystis, 280 Methylosinus, 280 Cytochrome o, Gluconobacter, 186–87

Cytochromes aerobic phototrophic bacteria, 571 Brucella, 367 dimorphic prosthecate bacteria, 74 Paracoccus, 243–44 Zymomonas, 211–12 Cytokines, Neisseria, 616

D Dalapon degradation, Achromobacter/Alcaligenes strains, 685 Dechloromonas, 17, 888 Dechlorosoma, 17 Dechlorosoma suillum, 888 Defensins, 664 Deformin production, Bartonella, 471–72, 477 Dehalogenation, Xanthobacter, 307 Dehalospirillum, 25 Dehalospirillum multivorans, 25 Delftia, 16, 723, 725 Delftia acidovorans, 723, 726 Deltaproteobacteria, 3–29 bdellovibrio group, 23 morphology of, 21 myxobacteria, 22–23 sulfate- and sulfur-reducing bacteria, 23–24 Denitrification Achromobacter/Alcaligenes strains, 680–82 Azoarcus, 885 Chromobacterium, 741 Paracoccus, 238, 240–45 phototrophic alphaproteobacteria, 55 Seliberia, 588 Thiobacillus, 817 Dental plaque, Neisseria, 603 1-Deoxynojirimycin production, Gluconobacter, 189 Dermonecrotic toxin (DNT), Bordetella, 658, 660–61 Derxia, 155, 751–56 applications of, 756 cultivation of, 752–53 habitat of, 751 identification of, 753–54 isolation of, 751–52 morphology of, 753–54, 754–55 motility of, 755 physiology and biochemistry of, 755–56 preservation of, 753 species description of, 754–55 Derxia gummosa, 751–56 Derxia indica, 751 Desulfacinum, 24 Desulfobacter, 23–24 Desulfobulbus, 24 Desulfofaba, 24 Desulfofrigus, 24 Desulfohalobium, 23 Desulfomicrobium, 23 Desulfomonile, 24

Index Desulfonatronum, 23 Desulfonema, 23 Desulforhabdus, 24 Desulfosarcina, 23 Desulfovibrio, 23 Desulfovirga, 24 Desulfurella, 23–24 Desulfuromonas, 23 Desulfuromusa, 23 Di- and Trisaccharide utilization Acidomonas, 179 Asaia, 180 Azospirillum, 123 Kozakia, 190 Zymomonas, 202–3, 208–11 Diagnosis Burkholderia, 852 gonorrhea, 630 of gonorrhea, 604, 606 melioidosis, 852 of meningococcal disease, 604, 631 pertussis, 652–53 Dialysis sac culture, Spirillum, 702 Diazosomes, 885 Dichelobacter, 20 Dichelobacter nodosus, 20 Dihydroxyacetone production, Gluconobacter, 189 3,6-Dihydroxyindoxazene, 741 N,N-Dimethylformamide degradation, Achromobacter/Alcaligenes strains, 686 Dimethylsulfide (DMS) production, Alcaligenes strains, 683 Dimorphic prosthecate bacteria, 72–86 cultivation of, 83–85 distribution of, 75–77 ecology and applications of, 85–86 enrichment and isolation of, 78–81 identification of, 81–83 morphology of, 81–82 rationale for clustering of, 72–75 2,4-Dinitrotoluene degradation, Achromobacter/Alcaligenes strains, 685–86 Disease. See also Animal disease; Human disease; Plant disease agricultural, 651 Agrobacterium, 91–110, 105–9 Bartonella, 467–69, 480–84 Brucella, 381–429 Burkholderia, 852–53 Chromobacterium, 739 Ehrlichia, 514–15 Eikenella corrodens, 843–44 gonorrhea, 629–31 Herbaspirillum, 143 meningitis, 629–32 Neorickettsia, 517 Orientia, 507 pertussis, 650–51 Proteobacteria, 11–12 Rickettsia, 500–504 Rickettsiales, 493 Wolbachia, 516, 555

Disseminated gonococcal infection, 631 D-lysine production, Comamonas, 732 DMS. See Dimethylsulfide production DNA content, Neisseria, 624–28 DNA-DNA hybridization Acetobacter, 176 Acetobacteraceae, 166 Acidomonas, 179 aerobic phototrophic bacteria, 571 ammonia-oxidizing bacteria, 780, 783–85, 789, 793, 795 Ancalomicrobium, 65 Asaia, 180 Azospirillum, 115–16, 123, 125–26, 130 Bartonella, 478 Brucella, 315, 331, 334, 337 Coxiella, 529 dimorphic prosthecate bacteria, 74 Gluconacetobacter, 181 Gluconobacter, 186 Herbaspirillum, 141–42 Kozakia, 190 Methylobacterium, 262 Methylomonas, 269 Phenylobacterium, 254–55 phototrophic alphaproteobacteria, 49–50 Prosthecomicrobium, 65 DNA-rDNA hybridization, Herbaspirillum, 141 DNA-rRNA hybridization Acetobacteraceae, 163, 166 Azospirillum, 115–16, 123 Methylobacterium, 263 Proteobacteria, 5–6 DNT. See Dermonecrotic toxin Dogs Bordetella, 651 Neisseria, 603 Simonsiella, 829, 834 Dolphin habitat, Brucella, 335 Donkeys Alysiella, 832 Burkholderia, 849 Simonsiella, 832 Doxycycline sensitivity Brucella, 429 Coxiella, 537 Ehrlichia, 514 Orientia, 507 Rickettsia, 500, 505–6 Duganella, 888

E Ear habitat Ochrobactrum, 748 Oligella, 691 Ecology aerobic phototrophic bacteria, 580 Agrobacterium, 105 Alphaproteobacteria, 9–15 Alysiella, 837–38

901

Anaplasma, 510 Azoarcus, 886–87 Azonexus, 888 Azospira, 888 Azospirillum, 116–20 Azovibrio, 888 Betaproteobacteria, 15–18 Comamonas, 731 dimorphic prosthecate bacteria, 85–86 Ehrlichia, 513–14 Gammaproteobacteria, 18–21 Herbaspirillum, 143–44 Leptothrix-Sphaerotilus group, 762–64 Neorickettsia, 517 Paracoccus, 237–39 Proteobacteria, 9–25 Rickettsia, 500 Simonsiella, 837–38 Wolbachia, 515–16, 553–55 Ecophysiology, Bordetella, 653–62 Ectothiorhodospiraceae, 18–19 Ehrlicheae, 460–61 Anaplasma, 460–61 Cowdria, 460 Ehrlichia, 460–61 Ehrlichia, 13, 460–61, 493–94, 512–15, 547 disease from, 514–15 ecology of, 513–14 epidemiology of, 514 genetics of, 512–13 habitats of, 512 identification of, 512 isolation of, 512 morphology of, 460 phylogeny of, 512 physiology of, 512 preservation of, 512 taxonomy of, 512 Ehrlichia bovis, 460 Ehrlichia canis, 460, 512–14 Ehrlichia chaffeensis, 460, 510, 512–14 Ehrlichia equi, 460–61 Ehrlichia ewingii, 460, 494, 512, 514 Ehrlichia muris, 460, 512 Ehrlichia phagocytophila, 460–61 Ehrlichia platys, 460 Ehrlichia risticii, 460–61 Ehrlichia ruminantium, 494, 512–15 Ehrlichia sennetsu, 460–61 Ehrlichiaceae, 13 Ehrlichioses Ehrlichia, 514–15 prevention of, 515 Eikenella, 18, 828 Eikenella corrodens, 840–45 antigenic structure of, 842 antimicrobial susceptibility of, 844 biochemical characteristics of, 842 differentiation of, 843 growth characterization of, 841–42 habitat of, 840 identification of, 842 isolation of, 840–42 media for selection of, 840–41

902

Index

microscopic morphology of, 842 pathogenicity of, 842–44 Elephants Alysiella, 832 Simonsiella, 832 Embden-Meyerhof-Parnas pathway aerobic phototrophic bacteria, 577 Coxiella, 534 Paracoccus, 245 Endocarditis Bartonella, 462, 480, 482–84 Coxiella, 537–39 Endoplasmic reticulum, Brucella, 397–400, 407–9 Endosymbiosis methanotrophs, 272 Phyllobacterium, 749 Enterobacter agglomerans, 731 Enterobacteriaceae, 19–20, 727, 843 Brenneria, 19 Buchnera, 19 Erwinia, 19 Escherichia, 19 Pectobacterium, 19 Photorhabdus, 19 Salmonella, 19 Shigella, 19 Xenorhabdus, 19 Yersinia, 19 Entner-Doudoroff pathway aerobic phototrophic bacteria, 576–77 Brucella, 363 Gluconobacter, 188 Paracoccus, 245 Zymomonas, 203 Environmental stress, Brucella, 367–69, 403 Enzymes Agrobacterium, 94–96 Bartonella, 473 Brucella, 362–63 Coxiella, 532 Phenylobacterium, 255 Eperythrozoon ovis, 493 Epidemiology Anaplasma, 510–11 Bartonella, 479–80 Brucella, 372–81 Burkholderia, 851–52 Coxiella, 536 Ehrlichia, 514 Neisseria, 628–30 Orientia, 507 Rickettsia, 500 Epididymitis, Neisseria, 630–31 Epoxide formation, Xanthobacter, 307–8 Epsilonproteobacteria, 3–29 Campylobacteraceae, 24–25 Helicobacter group, 25 morphology of, 24 Equine habitat Burkholderia, 849 Comamonas, 726 Ehrlichia, 460 Taylorella equigenitalis, 689–90

Erwinia, 19 Erythritol pathway, Brucella, 364–67 Erythrobacter, 14, 562, 565, 570–72 Erythrobacter litoralis, 569, 573, 578, 580 Erythrobacter longus, 565, 570–71, 575–77 Erythrocyte parasitism Anaplasma, 508–9, 511 Anaplasmataceae, 493 Bartonella, 472–78, 484 Erythromicrobium, 14, 562, 565, 570–72, 576–77 Erythromicrobium ezovicum, 569–70, 572–73, 578 Erythromicrobium hydrolyticum, 569–71, 575–76, 578 Erythromicrobium ramosum, 570, 573, 575, 578, 580 Erythromonas, 562, 565, 571–72 Erythromonas ursincola, 565, 569, 573, 578 Erythromycin resistance Neisseria, 625 Xanthobacter, 303 Erythromycin sensitivity Coxiella, 537 Xanthobacter, 303 Escherichia, 19, 650 Escherichia coli, 19, 28, 361, 613, 667–68, 864 Estuarine environment aerobic phototrophic bacteria, 566 dimorphic prosthecate bacteria, 76 Methylobacter, 278 Seliberia, 585 Thiobacillus, 815 Ethanesulfonate degradation, Achromobacter/Alcaligenes strains, 684–85 Ethanol fermentation, Zymomonas, 203, 207–9, 212–14 Ethanol oxidation Acetobacter, 176–77 Acetobacteraceae, 163 Acidomonas, 179 Asaia, 179–80 Erythromicrobium, 576 Gluconacetobacter, 180–81, 183 Kozakia, 190 Ethanol tolerance, Zymomonas, 208–9 Eukaryotic cell prokaryotic endosymbionts of, 25–28, 463 proteobacteria associated with, 318–19 Rickettsia, 496 Rickettsiales, 457 Wolbachia, 547 Evolution Bordetella, 650 Simonsiella, 830–31 Exopolysaccharide, Agrobacterium, 97–99 Exospores, Methylosinus, 282 Eye habitat, Ochrobacterum, 748

F False branching Leptothrix discophora, 769 Leptothrix mobilis, 769 Leptothrix-Sphaerotilus group, 759 Sphaerotilus, 773 Fatty acid utilization aerobic phototrophic bacteria, 576 phototrophic alphaproteobacteria, 55–56 Fatty acids Acetobacteraceae, 163 Achromobacter, 676 Acidomonas, 179 aerobic phototrophic bacteria, 571 Alcaligenes, 676 Alphaproteobacteria, 13 Alysiella, 837 Bartonella, 470–71 Brucella, 316, 320, 350–51 Coxiella, 532 Kerstersia, 676 methanotrophs, 271–72 Methylobacter, 278 Methylocaldum, 277 Methylococcus, 275 Methylocystis, 280 Methylomicrobium, 279 Methylomonas, 277–78 Methylosinus, 281 Methylosphaera, 279 nomenclature of, 275 Oligella, 690 Paracoccus, 240–43 Pelistega, 690 Pigmentiphaga, 676 Proteobacteria, 7 Simonsiella, 837 Taylorella, 690 Xanthobacter, 303–4 Feces Achromobacter/Alcaligenes strains, 687 Alcaligenes, 677 Comamonas, 726 Ochrobactrum, 748 Feline habitat Bartonella, 462, 469, 479 Coxiella, 536 Simonsiella, 829, 834 Ferric hydroxide, LeptothrixSphaerotilus group, 759 Ferromanganese oxides, LeptothrixSphaerotilus group, 759 Filament Alysiella, 833–35 Simonsiella, 828, 833–35 Filamentous bacteria, 758–75 Alysiella, 828–38 Simonsiella, 828–38 Filamentous hemagglutinin, 663, 667 Bordetella, 656–57 Fimbriae, 667 Bordetella, 657, 665 Fish habitat Coxiella, 530

Index dimorphic prosthecate bacteria, 77 Fission, dimorphic prosthecate bacteria, 73 Flagella Acidomonas, 179 aerobic phototrophic bacteria, 571–72 Agrobacterium, 96 Azospirillum, 123, 125, 127 Bartonella, 462, 467, 470–71, 474–75, 477 bdellovibrio group, 23 Beijerinckia, 156–58 Burkholderia, 856 dimorphic prosthecate bacteria, 72–73, 81, 83 Epsilonproteobacteria, 24 Gluconacetobacter, 180 Gluconobacter, 185 Methylobacter, 278 Methylobacterium, 261 Methylomicrobium, 278 Methylomonas, 277 Proteobacteria, 3 Xanthobacter, 296, 301 Flavobacterium capsulatum, 742 Flour, Acetobacteraceae, 174 Flowers Acetobacter, 176 Acetobacteraceae, 172–73 Asaia, 179–80 Gluconacetobacter, 181 Gluconobacter, 186 Fluorobenzoate degradation, Achromobacter/Alcaligenes strains, 685 Fluoroquinolone sensitivity Brucella, 428 Coxiella, 537 Food, Janthinobacterium, 742 Formaldehyde production, methanotrophs, 266 Formate production, phototrophic alphaproteobacteria, 55 Formate utilization, phototrophic alphaproteobacteria, 56 Francisella, 20, 461 Francisella tularensis, 20 Frateuria, 21 Freshwater environment aerobic phototrophic bacteria, 562, 566 ammonia-oxidizing bacteria, 783–86, 804 Ancalomicrobium, 65–66 Aquaspirillum, 711 Azospirillum, 122 bdellovibrio group, 23 dimorphic prosthecate bacteria, 76, 83 Leptothrix, 764–65 Leptothrix cholodnii, 768 Leptothrix discophora, 769 Leptothrix lopholea, 768 Leptothrix mobilis, 769 Leptothrix pseudoochracea, 768 methanotrophs, 270–72 Methylobacterium, 258–59

nitrite-oxidizing bacteria, 863–66 phototrophic alphaproteobacteria, 44–45 phototrophic betaproteobacteria, 596 Prosthecomicrobium, 65–66 Seliberia, 585–86 Skermanella, 126 Sphaerotilus, 772 Spirillum, 701 Thiobacillus, 813, 815 Thiovulum, 25 Xanthobacter, 295 Fruit tree habitat, Agrobacterium, 92 Fruits Acetobacter, 176 Acetobacteraceae, 172–73 Gluconacetobacter, 181 Gluconobacter, 186

G Gallionella ferruginea, 17, 770 Gammaproteobacteria, 3–29 Aeromonadaceae, 19–20 Alteromonadaceae, 19–20 ammonia-oxidizing bacteria, 786 Cardiobacteriaceae, 20 Chromatiaceae, 18–19 Ectothiorhodospiraceae, 18–19 Enterobacteriaceae, 19–20 Halomonadaceae, 20 Legionellaceae, 21 Methylococcaceae, 20 Moraxellaceae, 20 orders and families of, 18 Pasteurellaceae, 19–20 Pseudomonadaceae, 20–21 Succinivibrionaceae, 19–20 sulfur oxidizing bacteria of, 812–14, 821–24 Vibrionaceae, 19–20 Gas vesicles Methylobacter, 278 Methylosphaera, 279 Prosthecomicrobium, 70 Gastrointestinal tract Eikenella corrodens, 840 Enterobacteriaceae, 19 G+C content Acetobacter, 177 Achromobacter, 676 aerobic phototrophic bacteria, 571 Alcaligenes, 676, 684 Alcaligenes defragrans, 677 Alcaligenes denitrificans, 689 Alcaligenes faecalis, 677, 683, 687–88 Alcaligenes faecalis S-6, 680 Alysiella, 832, 837 ammonia-oxidizing bacteria, 784–86, 795 Anaplasma, 509 Aquaspirillum, 710–11, 713 Azoarcus, 877, 879–85 Azonexus, 889 Azospira, 889 Azospirillum, 116

903

Azovibrio, 889 Bartonella, 478 Beijerinckia, 158–59 Brucella, 320, 354, 372 Chromobacterium, 738 Comamonas, 728 Coxiella, 535 Derxia, 755 dimorphic prosthecate bacteria, 74 Eikenella corrodens, 840 Gluconacetobacter, 183 Herbaspirillum, 142 Janthinobacterium, 741 Kerstersia, 676 Leptothrix, 766 Leptothrix-Sphaerotilus group, 761 Methylobacter, 278 Methylocaldum, 277 Methylococcus, 275 Methylocystis, 280 Methylomicrobium, 279 Methylomonas, 277 Methylosinus, 281 Methylosphaera, 279 Oligella, 675, 690–91 Paracoccus, 234, 240–43 Pelistega, 675, 690 Phenylobacterium, 254 phototrophic alphaproteobacteria, 50 Pigmentiphaga, 676 proteobacteria, 28 Rickettsia, 498 Simonsiella, 832, 837 Spirillum, 704–5 Taylorella, 675, 689–90 Xanthobacter, 290 Gelatin hydrolysis, phototrophic betaproteobacteria, 599 Gemmobacter, 14 Gene expression, Brucella, 371–72 Gene structure, Bartonella, 478–79 Gene transfer, Zymomonas, 212 Genetic engineering, Agrobacterium, 109–10 Genetic exchange Bartonella, 479 Brucella, 372 Genetics Acetobacter, 177–78 Acidomonas, 179 aerobic phototrophic bacteria, 578–80 Agrobacterium, 96–105 Anaplasma, 509–10 Azoarcus, 886 Azonexus, 890 Azospira, 890 Azospirillum, 126–32 Azovibrio, 890 Bartonella, 478–79 Brucella, 327–28, 330, 332–33, 368–72 clonal and reticulate, 332–33 Comamonas, 729–31 Coxiella, 535 Ehrlichia, 512–13

904

Index

Gluconacetobacter, 184–85 Gluconobacter, 187–88 methanotrophs, 281–84 Neisseria, 624–28 Neorickettsia, 516–17 Orientia, 506–7 Rickettsia, 498–500 selective silencing of, 650 Xanthobacter, 306–7 Zymomonas, 212–14 Genitourinary tract Eikenella corrodens, 840 Ochrobactrum, 748 Oligella, 691 Genome Bartonella, 478 Brucella, 321–24, 368 Coxiella, 535 Rickettsia, 498 Wolbachia, 552 Gentamycin sensitivity, Methylobacterium, 262 Geobacter, 24 Glanders, 848, 849, 851–52, 853, 856 Gluconacetobacter, 13, 163–66, 180–85 applications of, 185 characteristics of, 168, 181, 186–87 classification of, 167, 180, 182 genetics of, 184–85 habitats of, 181 identification of, 181 isolation, cultivation and preservation of, 181 taxonomy of, 180–81 Gluconacetobacter azotocaptans, 173, 181, 184 Gluconacetobacter diazotrophicus, 119, 126, 173, 180–81, 184–85 Gluconacetobacter entanii, 172, 175, 181, 183 Gluconacetobacter europaeus, 175, 180–81, 183, 185 Gluconacetobacter hansenii, 173, 180–81, 183 Gluconacetobacter intermedius, 174, 181, 183–85 Gluconacetobacter johannae, 173, 181, 184 Gluconacetobacter liquefaciens, 169, 180–81 Gluconacetobacter oboediens, 181, 183 Gluconacetobacter sacchari, 173 Gluconacetobacter xylinus, 169, 171–74, 180, 183–84 Gluconacetobacter xylinus subsp. sucrofermentans, 183 Gluconate production Acidomonas, 179 Gluconacetobacter, 185 Gluconobacter, 188 Kozakia, 190 Gluconeogenesis, Coxiella, 534 Gluconobacter, 13, 151, 163–66, 168–70, 174, 185–89 applications of, 188–89 characteristics of, 168 classification of, 167

genetics of, 187–88 habitats of, 186 identification of, 186 isolation and cultivation of, 186 taxonomy of, 185–86 Gluconobacter asaii, 186 Gluconobacter cerinus, 186 Gluconobacter frateurii, 172–73, 186 Gluconobacter oxydans, 166, 169, 172–74, 185–86, 188–89 Glucose utilization Acidomonas, 179 Gluconacetobacter, 181 Gluconobacter, 186–87 Paracoccus, 245 Glutamate dehydrogenase, phototrophic betaproteobacteria, 598 Glycerol utilization Acidomonas, 179 Brucella, 363–64 Glycolytic pathway, Brucella, 364 Glycoproteins, Ehrlichia, 513 Glycosphingolipids, Sphingomonadaceae, 14 Goat habitat. See Caprine habitat Gold solubilization, Chromobacterium, 741 Gonococcal arthritis, Neisseria, 631 Gonococcus. See Neisseria gonorrhoeae Gonorrhea, 603, 630–31 antibiotic treatment of, 632 complications of, 630–31 diagnosis of, 604, 606, 630 disease incidence of, 629 prevention of, 631 Grahamella, 493. See also Bartonella Grape habitats Acetobacteraceae, 168–69 Agrobacterium, 91–92, 105–6, 109 Gluconobacter, 186 Grass roots, Azoarcus, 873, 875 Guinea pig habitat. See Rodent habitat

H Haemobartonella, 473 Haemophilus, 19, 841–42, 845 Haemophilus aphrophilus, 844–45 Haemophilus equigenitalis, 689 Haemophilus influenzae, 19, 28 Haemophilus parainfluenzae, 625 Haemophilus paraphrophilus, 844 Hairy root disease, Agrobacterium, 91, 105–6 Halomonadaceae, 20 Carnimonas, 20 Chromohalobacter, 20 Halomonas, 20 Zymobacter, 20 Halomonas, 20 Halothiobacillus, 18 Halothiobacillus neapolitanus, 816 Hazardous waste degradation, Xanthobacter, 308–9 Heartwater disease, Ehrlichia, 515

Heat shock proteins Agrobacterium, 101 Brucella, 403–4 Heavy metals detection of, Spirillum, 707–8 nitrite-oxidizing bacteria, 869 resistance to Achromobacter/Alcaligenes strains, 686 Comamonas, 731 Helical bacteria Aquaspirillum, 710–21 Spirillum, 701–8 Helicobacter bovis, 25 Helicobacter group, 25 Helicobacter, 25 Nautilia, 25 Thiomicrospira denitrificans, 25 Thiovulum, 25 Wolinella, 25 Helicobacter pylori, 24–25 Helicobacter suis, 25 Heme, Neisseria binding of, 615–16 Hemobartonella felis, 493 Hemobartonella muris, 493 Hemoglobin, Neisseria binding of, 615–16 Hemolysin production, Bartonella, 471–72 Hemolysis Bordetella, 660 Burkholderia, 854 Simonsiella, 835 Heparan sulfate proteoglycan (HSPG) receptors, 611 Herbaspirillum, 17, 141–47 applications of, 147 characteristics of, 145 habitats and ecology of, 143–44 history of, 141 identification of, 145–46 isolation of, 144 morphology of, 145 phylogenetic tree of, 142 preservation of, 144–45 taxonomy of, 141–43 Herbaspirillum chlorophenolicum, 141, 143–45 Herbaspirillum frisingense, 141–44, 146–47 Herbaspirillum lusitanum, 141, 143–44, 146 Herbaspirillum rubrisubalbicans, 141, 143–44, 146 Herbaspirillum seropedicae, 141–44, 146–47 Herbicide degradation, Phenylobacterium, 250 Hippea, 23 Hippopotami Alysiella, 832 Simonsiella, 832 Hirschia, 14 Holdfasts Leptothrix discophora, 768 Leptothrix lopholea, 768 Leptothrix-Sphaerotilus group, 759, 762 Sphaerotilus, 773

Index Holospora Caedibacter, 462 Holospora, 462 Holospora, 13, 462 Holospora obtusa, 462 Honey habitat, Zymomonas, 201 Hopanoids, Zymomonas, 208–9 Horizontal transmission Bordetella, 650 Simonsiella, 828 Horse habitat. See Equine habitat Horse manure, Aquaspirillum, 711 Hospital environment Burkholderia, 850, 851 Comamonas, 726 Methylobacterium, 258–59 Ochrobactrum, 748 Hot spring environment aerobic phototrophic bacteria, 565 phototrophic alphaproteobacteria, 45 Thiobacillus, 815 Human disease Alphaproteobacteria, 14–15 Anaplasma, 511 Anaplasmataceae, 493 Bartonella, 462, 467–69, 479 Betaproteobacteria, 16 Bilophila, 23 Brucella, 315, 335–38, 372–429 Campylobacteraceae, 24–25 Cardiobacteriaceae, 20 Chlamydia, 463 Coxiella, 531, 536 Ehrlichia, 460–61, 514–15 Enterobacteriaceae, 19 Francisella, 20 Gammaproteobacteria, 18 Legionella, 21 Neisseriaceae, 18 Orientia, 459–60 Pasteurellaceae, 19 Proteobacteria, 11 Rickettsia, 456–60 Human granulocytic ehrlichiosis, Ehrlichia, 460–61 Human habitats Achromobacter, 678 Alcaligenes, 677 Alysiella, 832, 833–34 Bordetella, 650–51 Eikenella corrodens, 840 Neisseria, 603 Ochrobactrum, 748 Simonsiella, 829, 832, 833–34 Human monocytotropic ehrlichiosis, Ehrlichia, 460, 514 Hydrazine degradation, Achromobacter/Alcaligenes strains, 686 Hydrogen oxidation Paracoccus, 238 Xanthobacter, 290 Hydrogen production, phototrophic alphaproteobacteria, 55, 58–59 Hydrogenophaga, 16, 723 Hydrogenophaga flava, 16 Hydrogenophilus, 17

Hydrogenovibrio, 20 Hydrophilic pathway, Brucella, 359–60 Hydrophobic pathway, Brucella, 360 Hydrothermal environment aerobic phototrophic bacteria, 566 dimorphic prosthecate bacteria, 75–76 Hydrothermal vents, Thiobacillus, 815 3-(3-Hydroxyphenyl)-propionate degradation, Comamonas, 730 Hydroxypyruvate reductase, Paracoccus, 235 Hyperinduction, Azoarcus, 885 Hypersaline environment ammonia-oxidizing bacteria, 804 phototrophic alphaproteobacteria, 45 Hyphomicrobiaceae, 15 Hyphomicrobium, 15 Pedomicrobium, 15 Rhodomicrobium, 15 Xanthobacter, 290, 294 Hyphomicrobium, 15, 65, 72–86, 225 Hyphomicrobium aestuarii, 75, 80 Hyphomicrobium facilis, 80 Hyphomicrobium vulgare, 80 Hyphomicrobium zavarzinii, 80 Hyphomonas, 14, 72–86 Hyphomonas hirschiana, 74 Hyphomonas neptunium, 85 Hyphomonas polymorpha, 75, 77 Hypocreales, 885

I Identification Acetobacter, 176 Achromobacter/Alcaligenes strains, 686–87 Acidomonas, 178–79 aerobic phototrophic bacteria, 568–71 Agrobacterium, 92–94 Alysiella, 835–37 Anaplasma, 508–9 Ancalomicrobium, 69–70 Aquaspirillum, 719–21 Asaia, 180 Azoarcus, 877–85 Azonexus, 888–89 Azospira, 888–89 Azospirillum, 123–26 Azovibrio, 888–89 Bartonella, 469–71 Beijerinckia, 154–57 Brucella, 325–26, 341–48 Burkholderia, 850–51 Chromobacterium, 740 Comamonas, 726–28 Coxiella, 531–33 Derxia, 753–54 dimorphic prosthecate bacteria, 81–83 Ehrlichia, 512 Eikenella corrodens, 842

905

Gluconacetobacter, 181 Gluconobacter, 186 Herbaspirillum, 145–46 Janthinobacterium, 742–43 Kozakia, 190 Leptothrix, 766–69 Leptothrix-Sphaerotilus group, 763–64 manganese-oxidizing bacteria, 228–29 methanotrophs, 274–81 Methylobacterium, 261–63 Methylococcaceae, 275–79 Methylocystaceae, 279–81 Neisseria, 604–6 Ochrobactrum, 748–49 Orientia, 506 Phenylobacterium, 252–56 phototrophic alphaproteobacteria, 49–54 phototrophic betaproteobacteria, 597–98 Phyllobacterium, 748–49 Prosthecomicrobium, 69–70 Rickettsia, 496 Seliberia, 588 Simonsiella, 835–37 Sphaerotilus, 773–74 Spirillum, 703 Thiobacillus, 820–21 Wolbachia, 551–52 Xanthobacter, 300–306 Zymomonas, 202–3 Ideonella, 758–59 Iminodisuccinate degradation, Achromobacter/Alcaligenes strains, 686 Immunity Bartonella, 484–85 to pertussis, 664–66, 667 Incidence gonorrhea, 629 meningitis, 629–30 Inclusion bodies, Xanthobacter, 301 Indole degradation, Achromobacter/Alcaligenes strains, 686 Industrial processes, LeptothrixSphaerotilus group, 763 Innate immune defenses, in respiratory tract, 662–64 Insertion sequence Brucella, 370–71 Coxiella, 535 Gluconacetobacter, 184 Interleukins, Brucella, 414–15 Intracellular state, Bordetella, 667–68 Intravenous tubing, Comamonas, 726 Iodobacter, 737–38 Iodobacter fluviatilis, 742 Iron chelation, Brucella, 405–6 Iron (Fe2+) oxidation Acidithiobacillus ferrooxidans, 823 Leptothrix, 770–71 Leptothrix ochracea, 766–67 Sphaerotilus, 773–74

906

Index

Iron seeps and springs, Leptothrix, 764–65 Iron uptake, Brucella, 362–63 Iron-binding proteins, Neisseria, 613–16 Isoenzyme typing, Neisseria, 628 Isolation Acetobacter, 176 Acetobacteraceae, 175 Acidithiobacillus ferrooxidans, 823 Acidithiobacillus thiooxidans, 824 Acidomonas, 178 aerobic phototrophic bacteria, 566–68 Agrobacterium, 92 Alysiella, 834–35 ammonia-oxidizing bacteria, 778–79 Anaplasma, 508 Ancalomicrobium, 66–68 Aquaspirillum, 711–19 Asaia, 180 Azoarcus, 876–77 Azonexus, 888 Azospira, 888 Azospirillum, 120–22 Azovibrio, 888 Bartonella, 469 Beijerinckia, 152–54 Bordetella, 651–53 Brucella, 337–41 Burkholderia, 850 Chromobacterium, 739–40 Comamonas, 726 Coxiella, 530–31 Derxia, 751–52 dimorphic prosthecate bacteria, 78–81 Ehrlichia, 512 Eikenella corrodens, 840–42 Gluconacetobacter, 181 Gluconobacter, 186 Herbaspirillum, 144 Janthinobacterium, 742 Kozakia, 190 Leptothrix, 765–66 manganese-oxidizing bacteria, 225–28 methanotrophs, 272–74 Methylobacterium, 259–61 Neisseria, 603–4 nitrite-oxidizing bacteria, 865–66 Ochrobactrum, 748 Orientia, 506 Paracoccus, 237–40 Phenylobacterium, 250–51 phototrophic alphaproteobacteria, 45–49 phototrophic betaproteobacteria, 596–97 Phyllobacterium, 748 Prosthecomicrobium, 66–68 Rickettsia, 496 Seliberia, 585–88 Simonsiella, 834–35 Sphaerotilus, 772–73 Spirillum, 701–3 Thiobacillus, 816–19

Wolbachia, 551 Xanthobacter, 297–99 Zymomonas, 201–2 Isoprene degradation, Alcaligenes strains, 682

J Janthinobacterium, 17, 720, 737, 741–43 applications of, 743 differentiating characteristics of, 738, 743 enrichment of, 742 habitat of, 741–42 identification of, 742–43 isolation of, 742 morphology of, 741 motility of, 741 preservation of, 743 Janthinobacterium lividum, 737, 740, 741–43, 751

K Kallar grass Azoarcus, 875, 887 Azospira, 888 Azovibrio, 888 Kanamycin sensitivity, Methylobacterium, 262 Kennel cough, 651 Kerstersia, 675–76, 678–89 physiology of, 679 Kerstersia gyiorum, 689 Ketogluconate production Gluconobacter, 185, 188 Kozakia, 190 Ketogulonicigenium, 14 Kidneys, Comamonas, 726 Kingella, 18, 828, 844–45 Kingella denitrificans, 845 Kingella indologenes, 845 Kingella kingae, 845 Klebsiella, 747, 749 Klebsiella oxytoca, 854 Klebsiella pneumoniae, 146 Knallgas bacteria, Xanthobacter, 290 Kombucha Acetobacteraceae, 174 Gluconacetobacter, 181 Kozakia, 163–66, 189–90 characteristics of, 168, 190 classification of, 167 habitats of, 190 identification of, 190 isolation and cultivation of, 190 taxonomy of, 189–90 Kozakia baliensis, 174, 180, 190 Krebs’ cycle, Paracoccus, 245

L Lactate oxidation, Acetobacter, 175–76 Lactate production, Beijerinckia, 158

Lactobacillus acidophilus, 607 Lactobacillus viridescens, 174 Lactoferrin, 613–14, 663–64 Neisseria, 614–15 Lantadene degradation, Alcaligenes strains, 682 Lawsonia, 23 Lawsonia intracellularis, 23 Leaf nodules, Phyllobacterium, 747, 749 Legionella, 21, 329 Legionella pneumophila, 21, 330, 499, 529 Legionellaceae, 21 Coxiella, 21 Legionella, 21 Rickettsiella, 21 Leptothrix, 16, 225, 764–72, 774–75 characteristics of, 766 habitat of, 764–65 identification of, 766–69 isolation of, 765–66 metal oxidation by, 770–71 physiology of, 770–72 sheath structure and composition in, 771–72 Leptothrix cholodnii, 759, 762, 765–66, 768–69 Leptothrix cholodnii SP-6, 771–72 Leptothrix discophora, 229–30, 593, 675, 762, 766, 768–70 Leptothrix discophora SS-1, 764, 770–71 Leptothrix lopholea, 759, 762, 766, 768 Leptothrix mobilis, 766, 769–70 preservation of, 769 Leptothrix ochracea, 758, 761–62, 766–67, 773 Leptothrix pseudoochracea, 766–68 Leptothrix-Sphaerotilus group, 758–75 common traits of, 759–61 distinguishing traits of, 761 ecology of, 762–64 morphology of, 761 phenotypic characteristics of, 759–62 phylogeny of, 758–62 physiology of, 761 reference strains of, 762 studying distribution and abundance of, 763–64 taxonomy of, 758–62 Leucothrix, 20 Lice habitat Bartonella, 479 Rickettsia, 500 Ligase chain reaction, for gonorrhea diagnosis, 606 Lincomycin sensitivity, Coxiella, 537 Lipid A Alphaproteobacteria, 7 Brucella, 330, 345, 352–56, 359–61 Coxiella, 532 Legionella, 330 Lipid-modified azurin, Neisseria, 613 Lipids, Brucella, 346, 349–51, 360–61

Index Lipoid bodies, Beijerinckia, 155–58, 160 Lipooligosaccharide Brucella, 353–54 Neisseria, 617–22 antigenic variation of, 617–19, 617–21 biosynthesis of, 620–21 expression of, 618 sialylation of, 618–19 structure of, 618, 619 Lipopolysaccharide Bartonella, 471 Bordetella, 661–62 Brucella, 316, 327–28, 331, 344–45, 349, 351–56, 358–59, 372, 391, 404–5, 410–11, 417–18, 422 Coxiella, 531–32 Phenylobacterium, 254–55 phototrophic alphaproteobacteria, 50 Xanthobacter, 302 Lipoprotein, Rickettsia, 500, 505 Lipoquinone Methylomicrobium, 279 Methylomonas, 278 Lithotrophy, nitrite-oxidizing bacteria, 869 Lophomonas, 723 Lophomonas alcaligenes, 723 LOS. See Lipooligosaccharide LPS receptors, 663 LPS-binding protein (LBP), 663 Lymphocytosis, Bordetella, 659 Lysosome inhibition, Brucella, 404– 5

M Macrolide sensitivity, dimorphic prosthecate bacteria, 74 Macrophages, Brucella, 388, 391–94, 398–401, 405–6, 409–15 Magnetospirillum magnetotacticum, 41 Maize habitat Azospirillum, 118–20, 133 Chromobacterium, 738–39 Herbaspirillum, 143 Malleobactin, Burkholderia, 854 Malonomonas rubra, 23–24 Malta fever, Brucella, 372 Manganese (Mn2+) oxidation Leptothrix, 766–71 Leptothrix-Sphaerotilus group, 759, 761–62 Manganese chemistry, 223–24 Manganese-oxidizing bacteria, 222–30 applications of, 229 controversy and perspectives of, 229–30 habitats of, 222–25 Hyphomicrobium, 225 identification of, 228–29 isolation and enrichment of, 225–28 Leptothrix, 225

Manganese-oxidizing factor, 770–71, 772 Mannheimia, 19 Marine environment aerobic phototrophic bacteria, 562, 566 Alcaligenes strains, 682–83 ammonia-oxidizing bacteria, 802–4 Ancalomicrobium, 65–66 bdellovibrio group, 23 dimorphic prosthecate bacteria, 76 Gammaproteobacteria, 19–20 nitrite-oxidizing bacteria, 863, 866 phototrophic alphaproteobacteria, 44–45 Prosthecomicrobium, 65–66 Psychrobacter, 20 Thiobacillus, 813, 815 Thiovulum, 25 Vibrionaceae, 19 Xanthobacter, 295 Marine mammal habitat, Brucella, 335 Meat, fermented, Acetobacteraceae, 175 Media aerobic phototrophic bacteria, 568 Agrobacterium, 93 Alcaligenes, 677 ammonia-oxidizing bacteria, 779–80 Ancalomicrobium, 68–69 Azospirillum, 120–22 Brucella, 339–43 dimorphic prosthecate bacteria, 83–85 Eikenella corrodens, 840–41 Herbaspirillum, 145 methanotrophs, 272–74 nitrite-oxidizing bacteria, 866 Paracoccus, 239–40 phototrophic alphaproteobacteria, 46–49 Prosthecomicrobium, 68–69 Spirillum, 702–3 Xanthobacter, 299 Melioidosis, 848, 850, 852, 855–56 Melittangium, 22 Menaquinones Deltaproteobacteria, 7–8, 21 Epsilonproteobacteria, 7–8, 24 Meningitis, 631–32. See also Meningococcal disease Meningococcal disease, 603, 631–32 antibiotic treatment of, 632 complications of, 631–32 diagnosis of, 604, 631 disease incidence of, 629–30 prevention of, 632 Meningococcus. See Neisseria meningitidis Mesorhizobium, 14–15 Metabolic engineering, Zymomonas, 212–14 Metabolism Bartonella, 471 Brucella, 349, 361–68

907

methanotrophs, 281 Neisseria, 607–8 Proteobacteria, 4–5 Zymomonas, 203–7 Metal working fluids, Comamonas, 726 Metallogenium, 225, 230 Methane monooxygenase methanotrophs, 266, 272–73, 282–84 Methylobacter, 278 Methylomicrobium, 279 Methylosphaera, 279 particulate, 282–83 soluble, 283 Methane oxidation methanotrophs, 266, 281, 282 Methylobacter, 278 Methylocystis, 280 Methylomicrobium, 279 Methylosphaera, 279 Methanococcus, 703 Methanol utilization Acidomonas, 179 dimorphic prosthecate bacteria, 83–84, 86 methanotrophs, 266, 281 Methylobacterium, 257–60 Methylocystis, 280 Methylomicrobium, 279 Methylosphaera, 279 phototrophic alphaproteobacteria, 56 Methanol yeast process, Acetobacteraceae, 174 Methanotrophs, 266–84. See also Methylococcaceae; Methylocystaceae applications of, 284 characteristics of, 267 habitats of, 270–72 detection, 271–72 endosymbionts, 272 identification of, 274–81 isolation and cultivation of, 272–74 phylogenetic tree of, 267–68 phylogeny of, 266–68 physiology and genetics of, 281–84 preservation of, 281 taxonomy of, 266, 268–70 types of, 266–68 Methylarcula, 14 Methylhydrazine degradation, Achromobacter/Alcaligenes strains, 686 Methylobacillus, 17 Methylobacter, 20, 268–69 identification of, 278–79 taxonomy of, 269 Methylobacter alcaliphilus, 269, 278 Methylobacter luteus, 269, 278 Methylobacter marinus, 269, 278 Methylobacter psychrophilus, 269, 271, 273, 278 Methylobacter whittenburyi, 269, 278 Methylobacterium, 15, 257–63 applications of, 263

908

Index

habitats of, 258–59 isolation and cultivation of, 259–61 morphology of, 261 preservation of, 263 taxonomy of, 257–58 Methylobacterium aminovorans, 258 Methylobacterium chloromethanicum, 258 Methylobacterium dichloromethanicum, 258 Methylobacterium ethanolicum, 258 Methylobacterium extorquens, 258 Methylobacterium fujisawaense, 258 Methylobacterium hypolimneticum, 258 Methylobacterium mesophilicum, 258 Methylobacterium organophilum, 258–59, 261 Methylobacterium rhodesianum, 258 Methylobacterium rhodinum, 258, 571 Methylobacterium thiocyanatum, 258 Methylobacterium zatmanii, 258 Methylocaldum, 266 identification of, 275–77 taxonomy of, 268 Methylocaldum gracile, 268, 277 Methylocaldum szegediense, 268, 277 Methylocaldum tepidum, 268, 277 Methylocapsa, 15 Methylococcaceae, 20, 266–84 characteristics of, 276 identification of, 275–79 Methylobacter, 20, 266 Methylocaldum, 266 Methylococcus, 20, 266 Methylomicrobium, 266 Methylomonas, 20, 266 Methylosphaera, 266 taxonomy of, 268–69 Methylococcus, 20, 266 identification of, 275 taxonomy of, 268 Methylococcus bovis. See Methylobacter luteus Methylococcus capsulatus, 268, 275, 283 Methylococcus chroococcus. See Methylobacter whittenburyi Methylococcus thermophilus, 268, 275 Methylococcus vinelandii. See Methylobacter whittenburyi Methylocystaceae, 266–84 identification of, 279–81 Methylocystis, 266 Methylosinus, 266 taxonomy of, 269–70 Methylocystis, 15, 266 identification of, 279–80 taxonomy of, 269–70 Methylocystis echinoides, 270, 280 Methylocystis minimus, 269–70 Methylocystis parvus, 269–70, 279–80, 282 Methylomicrobium, 266, 269 identification of, 278–79

taxonomy of, 269 Methylomicrobium agile, 269, 279 Methylomicrobium album, 269, 283 Methylomicrobium pelagicum, 269, 271, 279 Methylomonas, 20, 266 identification of, 277–78 taxonomy of, 268–69 Methylomonas aurantiaca, 277 Methylomonas flagellata, 269 Methylomonas fodinarum, 277 Methylomonas margaritae, 269 Methylomonas methanica, 268–69, 277 Methylomonas methaninitrificans, 269 Methylomonas methanooxidans, 269 Methylomonas pelagica. See Methylomicrobium Methylophaga, 20, 266 Methylophilus, 17, 269 Methylosinus, 266, 269 identification of, 280–81 taxonomy of, 270 Methylosinus sporium, 270, 280–81 Methylosinus trichosporium, 270, 280–84 Methylosphaera, 266 identification of, 279 taxonomy of, 269 Methylosphaera hansonii, 269, 271–73, 279, 283–84 Methylovibrio soehngenii, 270 Methylovorus, 17 Mice habitat. See Rodent habitat Micrococcus denitrificans. See Paracoccus denitrificans Micrococcus radiodurans, 688 Miglitol production, Gluconobacter, 189 Milk habitat, Comamonas, 726 Mitochondria Alphaproteobacteria, 3, 7, 13, 25–28 as prokaryotic endosymbiont, 463–64 Rickettsia, 495 MOF. See Manganese-oxidizing factor Molybdenum requirement, Beijerinckia, 160 Monkeys Alysiella, 832 Chromobacterium, 739 Simonsiella, 832 Monoclonal antibody Neisseria, 612, 618–19 nitrite-oxidizing bacteria, 863–65 Monosaccharide utilization Acidomonas, 179 aerobic phototrophic bacteria, 576 Asaia, 180 Azospirillum, 115, 123, 127 Brucella, 362–64, 367 Gluconacetobacter, 181 Gluconobacter, 186–87 Herbaspirillum, 145–46 Kozakia, 190 Paracoccus, 240–43

phototrophic alphaproteobacteria, 55–56 Xanthobacter, 290–91, 305 Zymomonas, 202–3, 208–11, 213 Moraxella, 20, 828, 835, 845 Moraxellaceae, 20 Acinetobacter, 20 Moraxella, 20 Psychrobacter, 20 Morphology aerobic phototrophic bacteria, 571 Agrobacterium, 94 Alphaproteobacteria, 9–15 Alysiella, 828, 830, 834–36 ammonia-oxidizing bacteria, 780, 783–86 Anaplasmataceae, 493 Ancalomicrobium, 65 Aquaspirillum, 719 Azoarcus, 877–85 Azonexus, 889 Azospira, 889 Azospirillum, 115, 121 Azovibrio, 889 Bartonella, 467, 470 bdellovibrio group, 23 Beijerinckia, 151–52, 154–59 Betaproteobacteria, 15–18 Brucella, 341–44, 349–61 Cardiobacterium hominis, 844 Chlamydiae, 463 Chromobacterium, 738 Comamonas, 726 Coxiella, 529 Deltaproteobacteria, 21 Derxia, 754–55 dimorphic prosthecate bacteria, 81–82 Ehrlichia, 460 Eikenella corrodens, 842 Epsilonproteobacteria, 24 Gammaproteobacteria, 18–21 Haemophilus paraphrophilus, 844 Herbaspirillum, 145 Janthinobacterium, 741 Leptothrix, 767–70 Leptothrix-Sphaerotilus group, 759, 761 Methylobacter, 278 Methylobacterium, 261 Methylocaldum, 275 Methylocystis, 279–80 Methylomicrobium, 278 Methylomonas, 277 Methylosinus, 280 Methylosphaera, 279 Myxobacteria, 22 Neisseria, 605–6 Paracoccus, 233–34 Phenylobacterium, 252–53 Prosthecomicrobium, 65, 70 Proteobacteria, 3, 9–25 Rhodocyclus, 597 Rickettsia, 457–59 Rubrivivax, 596 Seliberia, 585, 587 Simonsiella, 828, 830, 833–36 Skermanella, 126 Sphaerotilus, 773–74

Index Spirillum, 703 Wolbachia, 515, 551 Xanthobacter, 290, 294–96, 300–301 Mosquito habitat, Wolbachia, 547, 550–51 Motility Acidomonas, 179 Agrobacterium, 96 Alysiella, 833–35, 833–37 Aquaspirillum, 719 Azoarcus, 877 Azospirillum, 121, 123, 125 Bartonella, 471, 474 Beijerinckia, 156–59 Bordetella, 650 Burkholderia, 850–51, 856 Chromobacterium, 738 Deltaproteobacteria, 21 Derxia, 755 dimorphic prosthecate bacteria, 72–73 Eikenella corrodens, 841 Epsilonproteobacteria, 24 Gluconacetobacter, 180 Gluconobacter, 185 Herbaspirillum, 145 Janthinobacterium, 741 Methylobacter, 278 Methylobacterium, 261 Methylocaldum, 275 Methylomicrobium, 278 Methylomonas, 277 Myxobacteria, 22–23 Neisseria, 605, 622–23 Neisseriaceae, 18 Ochrobactrum, 748–49 Phyllobacterium, 748–49 Prosthecomicrobium, 70 Proteobacteria, 3 Simonsiella, 833–35, 833–37 Sphaerotilus, 773 Spirillum, 703, 707–8 Mottled stripe disease, Herbaspirillum, 143 Mouse habitat. See Rodent habitat Mouth. See Oral cavity Mtases, Neisseria, 626–27 Mucociliary defenses, Bordetella, 658 Mucosal surfaces Alysiella, 833 Comamonas, 726 Eikenella corrodens, 840 Neisseria, 603 Simonsiella, 833 Mule, Burkholderia, 849 Murine typhus, Rickettsia, 503–4 Mussels, Aquaspirillum, 711 Mutagenesis, Zymomonas, 212 Mycobacterium, 320 Mycobacterium flavum, 151 Mycobacterium tuberculosis, 371 Mycoplana, 747 Mycoplana rubra, 257 Myxobacteria, 22–23 morphology of, 22 Myxococcales, 22 Myxococcaceae, 22

Myxococcales, 22 Archangiaceae, 22 Cystobacteraceae, 22 Myxococcaceae, 22 Polyangiaceae, 22 Myxococcus, 22 Myxococcus xanthus, 28 Myxospores, Myxobacteria, 22

N Nalidixic acid resistance, Methylobacterium, 262 Nannocystis, 22 Naphthalene degradation, Comamonas, 729 Nata Acetobacter, 176 Acetobacteraceae, 173 Gluconacetobacter, 181 Native hapten polysaccharide, Brucella, 354, 356, 388–89, 391, 403 Nautilia, 25 Neisseria, 18, 602–32, 720, 828 bacteriophages of, 626 cellular structures of, 608–24 characteristics of, 605–6 disease from, 630–32 gonorrhea, 630–31 meningitis, 631–32 epidemiology of, 628–30 genetics of, 624–28 habitats of, 603 identification of, 604–6 isolation of, 603–4 metabolism of, 607–8 morphology of, 605–6 phylogeny of, 602 physiology of, 607–24 preservation of, 607 proteins of, 608–17 restriction and modification systems of, 626–27 species of, 602 strain typing of, 628–29 transformation of, 625–26 Neisseria animalis, 602–3, 609 Neisseria canis, 602–3, 609 Neisseria cinerea, 602–3, 606–9, 613, 622, 624–25 Neisseria denitrificans, 602, 609 Neisseria dentiae, 602 Neisseria elongata, 602, 605, 608 Neisseria flava, 608–9, 613 Neisseria flavescens, 602–3, 608–9 Neisseria gonorrhoeae, 18, 602–3, 606–30, 632 isolation of, 604 morphology of, 605 Neisseria gonorrhoeae 1291, 620 Neisseria gonorrhoeae FA19, 620 Neisseria lactamica, 602–3, 606–7, 609, 613, 622, 624–25 Neisseria macacae, 602–3 Neisseria meningitidis, 18, 602–3, 606–17, 621–32 isolation of, 604

909

Neisseria mucosa, 602–3, 606, 608, 613, 624–25 Neisseria perflava, 603, 608, 613, 624, 626 Neisseria polysaccharea, 602, 606–7, 609, 624 Neisseria sicca, 602–3, 606, 608–9, 613, 624–25 Neisseria subflava, 602–3, 606, 608, 613, 625 Neisseria weaveri, 602 Neisseriaceae, 18, 737, 828, 837 Alysiella, 18 Chromobacterium, 18 Eikenella, 18 Kingella, 18 Neisseria, 18 Simonsiella, 18 Nematodes, Alcaligenes, 677 Neorickettsia, 13, 460–61, 493, 516–17, 547 disease from, 517 ecology of, 517 genetics of, 516–17 habitats of, 516 physiology of, 517 taxonomy of, 516 Neorickettsia helminthoeca, 13, 461, 516 Neorickettsia, Neorickettsia, 460–61 Neorickettsia risticii, 516–17 Neorickettsia sennetsu, 516 Neptunomonas, 21 Neuroretinitis, Bartonella, 462 Neutrophils, 662–63, 664, 667–68 Brucella, 394 Nickel resistance Achromobacter/Alcaligenes strains, 686 Comamonas, 731 Nickel uptake, Brucella, 363 Nitrate reduction Azospirillum, 127 Brucella, 345, 367 Comamonas, 728 phototrophic alphaproteobacteria, 55 Thiobacillus, 817 Nitrates, manufacture of, 861 Nitric oxide reductase, Achromobacter/Alcaligenes strains, 680–81 Nitrilase Achromobacter/Alcaligenes strains, 687–88 Comamonas, 731 Nitrite oxidation, 861, 862 Nitrite oxidoreductase (NOR), 862, 864–65 Nitrite reductases, Achromobacter/Alcaligenes strains, 680–81 Nitrite reduction, Neisseria, 607 Nitrite-oxidizing bacteria, 861–69 biodegradation by, 868–69 biofilms of, 867 dominant members of, 863–65 enumeration of, 867–68 habitat of, 866–67

Prokaryotes (2006) 5:547–561 DOI: 10.1007/0-387-30745-1_22

CHAPTER 3.1.21 ehT

suneG

a i hcab l W o

The Genus Wolbachia MARKUS RIEGLER AND SCOTT L. O’NEILL

Introduction Numerous invertebrate species form long lasting symbioses with bacteria (Buchner, 1949; Buchner, 1965). One of the most common of these bacterial symbionts is Wolbachia pipientis, which has been estimated to infect anywhere from 15–75% of all insect species (Werren et al., 1995a; West et al., 1998; Jeyaprakash and Hoy, 2000; Werren and Windsor, 2000) as well as many species of arachnids, terrestrial crustaceans and filarial nematodes (O’Neill et al., 1997a; Bandi et al., 1998). In most arthropod associations, Wolbachia act as reproductive parasites manipulating the reproduction of their hosts to enhance their own vertical transmission. There appears to be little direct fitness cost to the infected host besides the costs arising from the reproductive manipulations. However instances have been reported where Wolbachia can be either deleterious (Min and Benzer, 1997; Bouchon et al., 1998) or beneficial (Girin and Boultreau, 1995; Stolk and Stouthamer, 1995; Wade and Chang, 1995; Vavre et al., 1999b; Dedeine et al., 2001) to their hosts. Wolbachia were first described as intracellular Rickettsia-like organisms (RLOs), infecting the gonad cells of the mosquito, Culex pipiens (Hertig and Wolbach, 1924), and were later named “Wolbachia pipientis” (Hertig, 1936). It was not until the work of Yen and Barr (Yen and Barr, 1971; Yen and Barr, 1973) that Wolbachia were implicated in causing crossing incompatibilities between different mosquito populations (Laven, 1951; Ghelelovitch, 1952). When polymerase chain reaction (PCR) diagnostics for Wolbachia became available, it became clear that this agent was both extremely widespread and also responsible for a range of different reproductive phenotypes in the different hosts it infected (O’Neill et al., 1992; Rousset et al., 1992; Stouthamer et al., 1993). The most common of these are cytoplasmic incompatibility, inducing parthenogenesis, overriding host sex-determination, and male-killing (O’Neill et al., 1997a). As of the time of this writing, more than 450 different Wolbachia strains with unique gene

sequences, different phenotypes, and infecting different hosts have been deposited in GenBank and the Wolbachia host database (http:// www.wolbachia.sols. uq.edu.au).

Phylogeny Wolbachia pipientis (Hertig, 1936) is the only species of the genus Wolbachia, family Anaplasmataceae, order Rickettsiales, class αproteobacteria (Dumler et al., 2001). Two other species, Wolbachia persica (Suitor and Weiss, 1961) and Wolbachia melophagi (Nöller, 1917; Philip, 1956), originally in the same genus have since been removed (Dumler et al., 2001). Wolbachia’s closest relatives in the Anaplasmataceae are the genera Anaplasma, Ehrlichia and Neorickettsia (Fig. 1). They are all obligate intracellular bacteria that reside in vacuoles of eukaryotic cells (Dumler et al., 2001). On the basis of the phylogeny of the 16S rRNA gene, the genus Wolbachia as currently defined is monophyletic (O’Neill et al., 1992; Bandi et al., 1998; Lo et al., 2002). Wolbachia have been divided into six supergroups A–F on the basis of 16S rRNA and ftsZ gene sequences; A, B, E and F are associated with arthropods (Vandekerckhove et al., 1999; Lo et al., 2002), and C and D are associated with filarial nematodes (Bandi et al., 1998; Lo et al., 2002). The phylogenetic relationship between these supergroups is currently not well resolved (Lo et al., 2002; Fig. 2). The validity of the A and B supergroups that infect insects has been confirmed by phylogenetic analysis of the heat shock operon groE (Masui et al., 1997), of the spacer 2 region between the 23S and the 5S rRNA coding genes (Van Meer et al., 1999) and of the surface protein gene wsp (Zhou et al., 1998; Fig. 3). There is evidence that homologous recombination between different Wolbachia strains occurs (Jiggins et al., 2001; Werren and Bartos, 2001) and thereby has the potential to confound the interpretation of phylogenetic data based on the sequences of single genes. The lack of congruency between phylogenetic trees of Wolbachia

548

M. Riegler and S.L. O’Neill

CHAPTER 3.1.21 U95927 U82963 Haemobartonella felis Haemobartonella muris

AE001345 Chlamydia trachomatis

AF016546 Eperythrozoon wenyonii 100

U88565 Eperythrozoon suis

100

0.1

AB00152 Omithoaoros moubata tick symbiote A

AB001519 Haernaphysalis fongicocnis tick symbiote A D84559 Coxiella sp.

X89110 Wolbachia melophagi

100

91

D38622 Ovienba tsutsugamushi

100

U12457 N. heliminthoeca AF036651 AF170727 E. rislicii AF170729 M21290 E. rislicii 100 AF036656 AF036656 E. rislicii AF036650 AF036659 AF036657 AF036655 E. rislicii AF036652 84 AF036648 AF036649 E. rislicii U34280 Ehruchia sp. SF agent AF036654 AF036653 E. rislicii AF037211 AF037210 M73219 E. sennetsu 64 M73225

100

U12463 100 Rickettsia typhi 63 100 97 D84558 L36217 Ricketrsia rickettsii

82

AB001522 Omithoaoros moubata tick symbiote B

76

JD1859 E. coli

U12457

Neorickettsia

98

AF059062 Ehrlichia sp. (abalone)

AF104680

Wolbachia

100

AF179630 from Folsomia candida AF035160 AJ010276 in Onchpcerca ochengi 749261 in Dirolilana immilis AF069068 AJ010275 in Brugia malayi 88 AJ012646 AF220604 U17059 L02882 in Muscidifurax uriraptor U83098 U83093 U83091 U83092 in Gryllus rubens 83 U83094 L02887 U80584 L02883 L02888 in Trichogrvamma deion L02884 U83097 U83090 64 U83095 in Gryllus integer U83095 U17050 U23709 81 X61768 in Culex pipiens AB025965

54 54

72

M21292 Wolbachia persica

Ehrlichia Anaplasma M60313 A. marginale AF283007 A. Centrale AF309865 72 AF311303 AF309867 A. marginale AF309868 AF309866 AF316944 A. Centrale AF309869 U03775 E. bovis AF316945 96 NKIT365866 A. ovis 99 AF057707 AF136714 AF169153 AF136712 U72879 Ehrlichia sp. P. leucopus U72878 ES242785 U02521 Ehrlichia sp. HGE agent U77389 U10873 ES242783 AF084907 AF093788 Ehrlichia sp. HGE agent AF012528 AF136713 AF241532 AF170728 AF036647 AF036646 E. equi in I. pacificus AF036645 ES242784 M73224 E. phagocytophila M73223 E. equi M73220 E. phagocytophila AF172167 AF172166 AF172165 AF172164 AF093789 AF156784 U54805 E. platys M82801 U52514 Ehrlichia sp. in A. americanum U27102 U27101 Ehrlichia sp.WTD 100 U27103 U27104

63

AF104680 Ehrlichia sp. Schotti variant AF1069758 99 U03777 U03776 Cowdria ruminardium X61659 X62432 U23503 96 AE147752 M73222 Ehrlichia chaffeensis U86665 U86664 U60476 U96436 M73227 Ehrlichia ewingii AB013008 AB013009 U15527 Ehrlichia muris AF162860 AF318946 Ehrlichia ovis 97 U54805 U26740 Ehrlichia caris 65 M73221 M73226 Ehrlichia caris

99

Fig. 1. Phylogenetic tree inferred from the small subunit (16S) rRNA gene sequences of Ehrlichia, Anaplasma, Neorickettsia and Wolbachia species, including 455 sites after removal of sites containing a gap in any sequence. The sequence from Chlamydia trachomatis (accession no AE001345 [www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=nucleotide=search=AE001345]) was used as an outgroup. Numbers above internal nodes indicate the percentage of 1000 bootstrap replicates that supported the branch. All bootstrap values are included for clades that were consistently observed using the phylogenetic methods applied (maximum parsimony, minimum evolution, maximum likelihood and majority-rule bootstrap analysis of neighborjoining trees). The maximum-likelihood tree is shown. Bars, estimated number of substitutions per site; the scales for the figure and insets are the same. From Dumler et al. (2001).

CHAPTER 3.1.21

The Genus Wolbachia

549

Onchocerca gutturosa [AJ010266]

Dirofilaria repens [AJO1273] Dirofilaria immitis [AJ010272]

Onchocerca ochengi [AJ010268]

0.05 substitutions/site

C 100 100

Brugia pahangi [AJ010270]

Supergroup Host range A, B various arthropods C, D filarial nematodes E springtail F termites

root 1

Brugia malayi [AJ010269]

D

root 3

100 100

Microcerotermes sp. [AJ292346]

87 1% NaCl NaCl catabolized conditions) (mol%)

Freshwater Bovine rumen

Humans and rodentsl Chickensu Antarctic soil

Marine

Habitat

a

Symbols: +, all species positive except where noted; −, all species negative except where noted; D, differs among species; BP, both poles; SP, single pole; Aer, aerobic; Micr, microaerophilic; ND, no data. b A. delicatum is mainly vibrioid. c A. delicatum has mainly a single flagellum at one pole; A. polymorphum has mainly a single flagellum at each pole. d A. putridiconchylium is catalase-negative or very weakly positive. e A. gracile, A. itersonii, and A. peregrinum can catabolize only a very restricted variety of sugars. f A. peregrinum and some strains of A. itersonii have nitrogenase activity. g Described by Grabovich et al., 1987, Microbiologiya 56:666–672. h May be microaerophilic on primary isolation. Although A. nitrofigilis is a microaerophilic nitrogen fixer, it can grow aerobically in complex media such as Brucella agar. i A butzleri is weakly catalase positive. j Some cells in Azospirillum cultures are straight rods. k Some Azospirillum species form numerous lateral flagella in addition to the polar flagellum when cultured on solid media. l All wild-type strains of Bdellovibrio upon initial isolation are dependent on intraperiplasmic growth in susceptible bacterial prey. Mutants capable of axenic growth (“prey-independent” strains) have been derived from the predacious strains, and some strains are facultative, i.e., capable of growth in the presence and absence of prey cells. m Chains of Campylobacter cells may have a helical appearance. C. rectus, C. showae, and C. gracilis are straight rods. n C. showae has a tuft of flagella at one pole. o H. trogontum and H. bilis are fusiform straight rods. H. bizzozeronii and H. felis cells are long helices. H. cholecystus cells are coccoid to short curved rods. p Some cells in cultures of Herbaspirillum are helical. q O. kriegii and O. jannaschii are straight rods. r O. pusillum has mainly a single flagellum at each pole. O. kriegii and O. jannaschii have a single flagellum at one pole. s Causes one form of rat-bite fever in humans. u Causes a diphtheroid stomatitis in chickens. t Produces acid fiomxylose but not from other sugars that have been tested; it is also distinguished by failing to grow at temperatures above 20°C. w Predacious on eukaryotic algae.

“Spirillumpulli” “Spinillum pleomorphum” Vampirovibrio Wolinella succinogenes

Pseudoalteromonas Curved undina “Spirillumminus” Helical

Cell shape

Cell Requires diameter host for Oxygen (µm) growth requirement

CHAPTER 3.2.5 The Genus Spirillum 705

706

N.R. Krieg

Fig. 1. Polar flagellar fascicles of S. volutans ATCC 19554 n by electron microscopy. Bar = 1 µm. (From Hylemon et al., 1973a.)

below the surface at a point where the respiratory rate of the cells matches the rate of diffusion of oxygen to the cells. As the cell numbers increase, the disk becomes denser and migrates toward the surface. After 48 h dense growth occurs just beneath the surface. Cultures grown in MPSS broth (no agar added) or on solid media for colony development should be incubated under an atmosphere of 6% O2 and 94% N2. A convenient way to do this is to use a polycarbonate jar such as those that are used routinely for cultivation of anaerobes, provided that it has a vent to allow evacuation. After placing the inoculated media in the jar and sealing it, the jar is evacuated to 0.286 of the normal atmospheric pressure. Nitrogen gas is then added until the atmospheric pressure is restored. Padgett et al. (1982) reported that cells inoculated into MPSS broth can grow under aerobic conditions if the medium is supplemented with either bovine superoxide dismutase (SOD) (4 units/ml), catalase (0.8 units/ml), potassium metabisulfite (0.02%), or norepinephrine (0.002%). Cultures in CHSS broth (no agar added) can grow aerobically (Bowdre and Krieg, 1974). Five-ml volumes contained in 20 × 125-mm loosely screw-capped tubes are inoculated with several drops of culture and incubated at 30°C in a slanted position. Growth occurs near the surface of the medium in 24 h. Transfers must be made daily, with several drops of each previous culture serving as inoculum for the next tube. Cultures usually die within 48 h, possibly due to the high pH (>8.0) that develops as the result of oxidation of the succinate. Bowdre et al. (1976) used a chemically defined liquid medium containing norepinephrine, which, like CHSS broth, supports growth of S. volutans under aerobic conditions. Growth of S. volutans on the surface of solid media (MPSS or CHSS broth solidified

CHAPTER 3.2.5

with 1.5% agar) is difficult but can be achieved by methods described by Padgett et al. (1982). The agar media must be supplemented with potassium metabisulfite (0.002%), catalase (230 units/ml), or SOD (30 units/ml). If catalase or SOD is used, it is added aseptically to the molten medium at 45°C just before dispensing into Petri dishes, whereas potassium metabisulfite is added prior to autoclaving. The medium is dispensed into Petri dishes (20 ml per dish) under dim illumination or red photographic safelight (red light does not cause photochemical generation of toxic forms of oxygen in the medium). The plates are allowed to dry in the dark at room temperature for 24 h. In dim illumination, 0.1-ml portions of appropriate dilutions of a broth culture of S. volutans are spread onto the agar surface with a glass rod. The plates are incubated for 5 days at 30°C in the dark in a sealed vessel containing 6– 12% O2 (the balance being N2). It is essential that the vessel be lined with moistened filter paper to maintain a high humidity. Even with these precautions, only 22–72% of the cells inoculated onto the plates develop into colonies. A method for obtaining reproducible colony counts was reported by Alban and Krieg (1996). The method uses a semisolid version of colony counting medium (CCM) as an overlay for a thicker layer of sterile medium. The medium contains pyruvate, which destroys hydrogen peroxide. Cells to be used for colony counts are grown in 5.0 ml of CCM broth contained in slanted 20 × 120-mm loosely screw-capped tubes incubated aerobically for 24 h at 30°C. Sterile semisolid medium (CCM plus 0.7% agar) is dispensed into Petri dishes (15 ml per dish) and allowed to gel for 30 min. A 0.1-ml volume of an appropriate dilution of the inoculum is added to 10 ml of semisolid CCM at 45°C and is poured onto the plates as an overlay. After this has gelled, the plates are incubated at 30°C in an atmosphere of 6% O2 and 94% N2 for 3–4 days.

Preservation Preservation of S. volutans can be accomplished by the following procedure (Pauley and Krieg, 1974): Cells from a turbid 24-hour-old broth culture are centrifuged at 3,500 × g and washed once in nutrient broth. The cells are then suspended by gentle agitation in a small amount of nutrient broth containing 10% dimethyl sulfoxide (DMSO). The resulting dense suspension is allowed to incubate for 30 min at room temperature and then 0.4-ml volumes are dispensed into plastic or glass vials suitable for liquid nitrogen preservation. The vials are sealed, frozen in a mixture of dry ice and alcohol, and then stored by submersion in liquid nitrogen. For recovery of

CHAPTER 3.2.5

the organisms, the vials are thawed in water at 30°C and transferred to semisolid MPSS or CHSS medium.

Physiology S. volutans has a strictly respiratory type of metabolism with oxygen as the terminal electron acceptor. It is microaerophilic in ordinary liquid media, such as nutrient broth (in which it grows poorly) and MPSS broth, and on solid media, such as CHSS agar supplemented with bisulfite. Also, it can grow aerobically in special liquid media such as CHSS broth or the chemically defined medium of Bowdre et al. (1976). Growth is inhibited by very low concentrations of H2O2 (0.29 µM or greater), and the organism is rapidly killed by concentrations of 10 µM or greater (Alban and Krieg, 1988). The microaerophilic nature of the organism has been attributed to the high susceptibility of the organism to toxic forms of oxygen which occur spontaneously in MPSS or CHSS broth under aerobic conditions, particularly when the media are illuminated (Padgett et al., 1982). Destruction of toxic forms of oxygen by media supplements such as SOD, catalase, potassium metabisulfite, or norepinephrine allows the organism to grow aerobically in broth (Padgett et al., 1982). However, even with these supplements, S. volutans can form colonies on MPSS, CHSS, or CCM agar only under microaerobic conditions (12% O2 or less). It will not grow even under microaerobic conditions unless the media supplements are present. Padgett and Krieg (1986) described two variant strains of S. volutans which, unlike the wild type, could grow in an air atmosphere (i.e., 21% O2) on CHSS agar containing potassium metabisulfite. Cell extracts of those aerotolerant variants exhibited greater o-dianisidine peroxidase activity than did the wild type, and this enzyme may be important for aerotolerance. The catalase-negative microaerophile Spirillum volutans is killed rapidly by levels of H2O2 greater than 10 µM. Alban and Krieg (1998) described a mutant isolated by single-step mutagenesis with diethyl sulfate that was able to survive and grow after exposure to 40 µM H2O2 and was effective in eliminating H2O2 added to the medium. Nevertheless, the mutant was no more tolerant to O2 than the wild type. The only apparent phenotypic difference between the wild type and the mutant was that the mutant had high NADH peroxidase activity (0.072 IU per mg protein) whereas the wild type had no detectable activity (90% of strains positive; −, >90% of strains negative; D, result varies among strains; and nd, no data available. From Etchebehere et al. (2001). b From Gumaelius et al. (2001). c As tested with auxanographic techniques using API systems. d From Tamaoka et al. (1987). e From Willems et al. (1989). a

can be added here that the presence of hydroxy putrescine is characteristic for the genera of the β-Proteobacteria. Table 3 gives features for the differentiation of Comamonas species.

Cultivation Comamonas strains can be grown on nutrient agar and many other commonly used media such as tryptic soy agar and Columbia agar with or without blood. They are able to use many organic acids and amino acids, but few sugars (De Vos et al., 1985). They can degrade a variety of complex aromatic compounds, steroids and many manmade complex organic compounds. Optimal growth temperature is 30°C.

Preservation Comamonas cultures can be maintained on nutrient agar slants at 4°C for up to 2 months. Lyophilization or storing in liquid nitrogen can be used for long-term preservation of strains on the condition that cryoprotectants are added. Generally, the use of standard procedures is recommended.

Physiology Comamonas strains have an aerobic respiratory type of metabolism. Most strains of C. terrigena and C. testosteroni are capable of nitrate reduc-

tion, but cannot reduce nitrites (Willems et al., 1991), while all strains of C. nitrativorans and C. denitrificans can reduce nitrate all the way to N2. Although Comamonas strains were identified in a study of denitrifying biofilms at a water treatment plant, they were thought not to be true members of the biofilm community, but to have washed in from previous stages (Lemmer et al., 1997). Other data, however, confirm the denitrification capacity of some members of the genus (Vanbrabant et al., 1993). In particular, it has been shown that Comamonas strain SGLY2 is capable of aerobic denitrification and nitrogen production without nitrite building up. This strain has been studied for use in mixed-culture aerated reactors (Patureau et al., 1997). Comamonas strains are able to degrade a wide variety of aromatic compounds. Comamonas strain JS765 has a 1,2-dioxygenase able to break down nitrobenzene to nitrohydrodiol which spontaneously decomposes to catechol and nitrite. Catechol is then broken down via the meta-cleavage pathway, the genes of which have been sequenced (Parales et al., 1997; Genetics). This pathway has been studied in comparison with similar pathways in Pseudomonas strains (He and Spain, 1999). Comamonas testosteroni strains that degrade polycyclic aromatic compounds such as phenanthrene, naphthalene and anthracene have been described and the genes involved Genetics have been characterized (Goyal and Zylstra, 1996). The mechanism for phenol tolerance was studied in the phenoldegrading C. tesosteroni strains P15 and E23 (Yap et al., 1999).

CHAPTER 3.2.7

The meta-cleavage pathway is used by C. testosteroni for the degradation of 4-chlorophenol and related compounds (Hollender et al., 1997). Comamonas testosteroni can degrade 4-toluene sulfonate, which is thought to be taken up by an inducible secondary proton symport system (Locher et al., 1993). The degradation of 4toluene sulfonate and 4-toluene carboxylate involves oxygenation of the side chain to 4sulfobenzoate and terephthalate, respectively, which are then oxidized to protocatechuate. Genes involved in these pathways have been sequenced and characterized (Junker et al., 1997b; Genetics). Terephthalate is broken down by a terephthalate dioxygenase system to (1R, 2S)-dihydroxy-3,5-cyclohexadiene-1,4-dicarboxylic acid which is further degraded to protocatechuate by (1R,2S)-dihydroxy-1,4dicarboxy-3,5-cyclohexadiene dehydrogenase (Oppenberg et al., 1995). The dioxygenase system is a Rieske [2Fe-2S] protein with two subunits α and β which are thought to form an α2β2 structure (Schlaefi et al., 1994). The sulfonate and sulfate ester degradation as a source of sulfur for growth of Gram-negative bacteria has recently been reviewed (Kertesz, 2000). Comamonas testosteroni I2 was shown to degrade recalcitrant components such as aniline and chloroaniline (Boon et al., 2001). Both components are among the most widely produced industrial amines commonly used for the production of polyurethanes, rubber, azo dyes, drugs, pesticides, etc. The presence of these toxic amines in natural environments is strictly regulated by the environmental protection agencies of the European Union (EU) and the United States. Testosterone can be used as sole carbon source by C. testosteroni and induces the expression of enzymes involved in catabolism of steroids and aromatic hydrocarbons and the repression of at least one amino acid degrading enzyme. It was suggested that steroids may play a regulatory role in the synthesis of catabolic enzymes during adaptive growth (Moebus et al., 1997). Several of the enzymes involved in the conversion and break down of steroids have been studied and their genes sequenced (Genetics). The structure and function of δ5-3-ketosteroid isomerase and 3-oxo-δ5-steroid isomerase, which catalyze the the conversion of δ5- to δ4-3-ketosteroids by intramolecular proton transfer, have been studied (Brothers et al., 1995; Zao et al., 1996, 1997). A bile acid 3-α-sulfate sulfohydrolase of C. testosteroni was purified and characterized (Tazuke et al., 1994). Comamonas strains enriched from wastewater from an aerated stabilization basin of a bleached kraft pulp mill were reported to degrade terpenes such as abietane and pimarane-type resin acids (Morgan and Wyndham, 1996).

Comamonas

729

Some Comamonas strains are able to degrade poly-3-hydroxybutyrate using an extracellular poly-3-hydroxybutyrate depolymerase, which can also hydrolyze poly(3-hydroxybutyrate-co3-hydroxyvalerate) and poly-3-hydroxyvalerate. The enzyme differs from other poly-3-hydroxybutyrate depolymerases in that it is insensitive to phenylmethylsulfonyl fluoride and hydrolyzes poly-3-hydroxybutyrate to 3-hydroxybutyrate monomers (Jendrossek et al., 1993). The encoding gene of a Comamonas sp. strain was cloned and partly sequenced, and the resulting partial protein was analyzed (Jendrossek et al., 1995). A very similar sequence was obtained for C. testosteroni strain YM1004 (Shinomiya et al., 1997). Comamonas sp. strain P37C, isolated from compost, was able to degrade a range of poly-3hydroxyalkanoates (Quinteros et al., 1999). The enzymes involved in the first steps of quinoline and 3-methyl quinoline degradation by C. testosteroni strain 63, quinoline 2-oxidoreductase and 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase, have been characterized. Quinoline 2oxidoreductase is a molybdo-iron flavoprotein and 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase is a single component protein (Schach et al., 1995). Comamonas testosteroni produces 5-aminolevulinic acid, a precursor to tetrapyrroles, from aminoacylated tRNA-Glu via a two-step pathway involving glutamyl-tRNA reductase and glutamine-1-semialdehyde-2,1-aminomutase (Hungerer et al., 1995). The biosynthesis of aromatic amino acids via several pathways has been extensively studied among pseudomonad bacteria. In C. testosteroni prephenate dehydrogenase and arogenate dehydrogenase are reactive with either nicotinamide adenine dinucleotide (NAD) or nicotinamide adenine dinucleotide phosphate (NADP). No arogenate dehydratase activity is present; prephenate dehydratase is present and its activity is stimulated by L-tyrosine (Byng et al., 1983).

Genetics The genes of C. testosteroni involved in the degradation of naphthalene and phenanthrene have been cloned and characterized. At least two different sets of phenanthrene degradation genes may be present among different Comamonas testosteroni strains. They are different from the naphthalene degradation genes (nha) of Pseudomonas putida (Goyal and Zylstra, 1996). The gene phtD for 4,5-dihydroxyphthalate decarboxylase, an enzyme involved in phthalate metabolism in C. testosteroni, has been identified and sequenced. The deduced amino acid sequence shows similarity to the pth5 genes of P.

730

A. Willems and P. De Vos

putida, and the sequence upstream of pthD shows a high similarity to pth1 in P. putida, where it is thought to be a positive regulator of the other pth genes. These findings suggest a common origin for the genes involved in the phthalate metabolism (Lee et al., 1994). The bphD gene of C. testosteroni strain B-356, encoding for 2-hydroxy-6-oxo (phenyl/chlorophenyl) hexa-2,4-dienoic acid hydrolase, was sequenced, and the gene product has a mechanism of action similar to that of classical lipases and hydrolases (Ahmad et al., 1995). In the same strain, biphenyl/chlorobiphenyl dioxygenase, the enzyme system involved in the first step of the degradation of biphenyl or chlorobiphenyl compounds, was characterized. The genes for the three components of this enzyme system have been sequenced: bphA and bphE encode the two subunits of a terminal Fe-S oxygenase, bphF encodes a ferredoxin, and bphG encodes a ferredoxin reductase and is not located near the bphAEF genes (Sylvestre et al., 1996b). Many of these enzymes have been characterized (Hurtubise et al., 1995; Hurtubise et al., 1996) and some have been used to produce enzyme chimeras by combination with oxygenase components from other bacteria (Chebrou et al., 1999). Genes for subsequent degradation steps have also been characterized: bphB encodes 2,3dihydro-2,3-dihydroxybiphenyl dehydrogenase (Sylvestre et al., 1996a) and bphC encodes 2,3dihydroxybiphenyl-1,2-dioxygenase which catalyzes meta-1,2 fission of the aromatic ring (Bergeron et al., 1994). Finally, it has been demonstrated (Hein et al., 1998) that in this strain two meta-cleavage dioxygenases coexist with different degrees of sensitivity towards intermediates of the biphenyl catabolic pathway and suggesting a complex regulation system. Genes involved in the degradation of catechol via the meta-cleavage pathway have been sequenced and amino acid sequences deduced for strain Comamonas sp. JS765: cdoE encodes catechol 2,3-dioxygenase, cdoT encodes a ferredoxin, and cdoR is a regulatory gene (Parales et al., 1997). A gene cluster encoding for the degradation of 3-(3-hydroxyphenyl)-propionate from C. testosteroni strain TA441 was cloned and sequenced. It contains a regulatory gene, mhpR, and the following six genes for catabolic enzymes: mhpA for a flavin-type hydroxylase, mhpB for an extradiol dioxygenase, mhpD for 2-keto-4-pentenoate hydratase, mhpF for an acylating acetaldehyde dehydrogenase, mhpE for 4-hydroxy-2-ketovalerate aldolase, and mhpC for a meta-cleavage hydrolase (Arai et al., 1999b). The same research group (Arai et al., 2000) studied the arrangement and the regulation of the meta pathway in the breakdown of phenol.

CHAPTER 3.2.7

The genes involved in the degradation of 4toluene sulfonate and 4-toluene carboxylate of C. testosteroni have been extensively studied. The first step of oxygenation of the side chain to 4-sulfobenzoate and terephthalate, respectively, requires three enzymes: a 4-toluenesulfonate methyl-monooxygenase system consisting of a reductase B and oxygenase M encoded by the tsaBM genes, a 4-sulfobenzyl alcohol dehydrogenase encoded by tsaC and a 4-sulfobenzaldehyde encoded by tsaD. These genes are coexpressed under the regulation of the product of tsaR. Some of the enzymes involved have been purified and partially characterized. The tsaMB oxygenase system is a class IA mononuclear iron oxidase with tsaM having a Rieske [2Fe-2S] center, and tsaC is a short-chain zinc-dependent dehydrogenase (Junker et al., 1996). The conversion of 4-sulfobenzoate to protocatechuate is catalyzed by a 4-sulfobenzate-3,4-dioxygenase system consisting of a reductase C and an oxygenase A, again with a Rieske [2Fe-2S] center, encoded by the genes psbAC (Junker et al., 1996). The genes tsaMBCD, tsaR, and psbAC are located on conjugative plasmids in some C. testosteroni strains (Junker and Cook, 1997a). Terephthalate is also broken down to protocatechuate, but genes involved have not yet been characterized. Protocatechuate is broken down via a meta-cleavage pathway by enzymes thought to be encoded on the chromosome (Junker and Cook, 1997a). The gene cluster encoding a phenol hydroxylase in the phenol-degrading C. testosteroni strain R5 (phcKLMNOP) and the regulatory gene phcR have been cloned and expressed in Pseudomonas aeruginosa, conferring the use of phenol as sole carbon source on this bacterium (Teramoto et al., 1999). A second gene cluster which can confer the ability to use phenol (aphKLMNOPQB and the corresponding regulatory genes aphR and aphS) was studied in C. testosteroni strain TA441 (Arai et al., 1999a). In C. testosteroni, the genes for several enzymes involved in the breakdown of steroid compounds have been studied. The gene for a 3-α-hydroxysteroid dehydrogenase has been sequenced and expressed through cloning, and the enzyme was characterized (Abalain et al., 1995; Oppermann and Maser, 1996a). Because of its similarity to ribosomal proteins L10 and L7/ 12, it was suggested that this enzyme may be formed by fusion of two ribosomal proteins (Baker, 1996). The gene for 3-ketosteroid-δ4-5-αdehydrogenase was sequenced and located downstream of a gene for 3-ketosteroid-δ1dehydrogenase. Although both enzymes are functionally similar, their genes are probably not derived from a common ancestor (Florin et al., 1996). A 3β/17β hydroxysteroid dehydrogenase

CHAPTER 3.2.7

was studied by mutagenic replacement within the active site to identify important residues (Oppermann et al., 1997). The steroid-inducible gene stdC of C. testosteroni was cloned and sequenced. It was homologous to open reading frames (ORFs) of unknown function in the polyhydroxyalkanoic acid cluster of several other bacteria and is transcribed as a monocistronic mRNA that is abundant in cells grown on steroid carbon sources (Cabrera et al., 1997). Recently, in a model for the regulation of the 3-αhydroxysteroid hydrogenase/carbonyl reductase (Xiong et al., 2001), it was proposed that the hydrogenase is under the control of two repressor proteins of which the binding to the promotor and the messenger RNA is de-repressed by an inducer such as testosterone. The gene for an aliphatic nitrilase from C. testosteroni was sequenced and overexpressed, and the enzyme was purified and its primary structure determined. A Cys163 residue is thought be essential in the active site (Levy-Schil et al., 1995). Several plasmids have been isolated from or transferred to Comamonas strains, and they may contribute to the catabolic versatility of these organisms. Comamonas testosteroni strains T-2 and PSB-4 and the type strain were reported to contain plasmids pTSA and pT2T, plasmid pPSB, and no plasmids, respectively (Junker and Cook, 1997a). Plasmid pTSA (85 kb) is a conjugative plasmid of the IncP1 group and carries the genes tsaMBCD, tsaR and psbAC, involved in the degradation of 4-toluene sulfonate to protocatechuate, and two copies of insertion sequence IS1071. Plasmid pPSB (85 kb) also is a conjugative plasmid; it carries only the psbAC gene as well as two copies of IS1071. Conjugative experiments with the type strain have suggested that the psb genes are located in a composite transposon (Junker and Cook, 1997a). Plasmid RP4::Tn4371 with genes for biphenyl and 4-chlorobiphenyl degradation was transferred from Enterobacter agglomerans to indigenous soil bacteria, including Comamonas strains, where these catabolic enzymes were successfully expressed (De Rore et al., 1994). Plasmid PR4::Mu3A, carrying chromosomal DNA fragments encoding for the use of biphenyl as a sole carbon source, were transferred to C. testosteroni and the enzymes were effectively expressed (Springael et al., 1996). Plasmids pACK5 and pACT72, including a replicon of the cryptic Acetobacter pasteurianus plasmid pAC1, were transferred to and successfully expressed in C. terrigena (Grones and Turna, 1995). Plasmid pE43 from Pseudomonas aeruginosa and plasmid pPC3 from Arthrobacter globiformis, carrying genes for dehalogenation of chlorobenzoates, permitted C. testosteroni strain VP44 to grow on

Comamonas

731

ortho- or para-biphenyls as sole carbon source (Hrywna et al., 1999). Plasmid PNB2 (Boon et al., 2001) has been isolated from Comamonas testosteroni I2, an aniline- and chloroaniline-degrading strain. So far, the chloroaniline capacity could not be demonstrated on the plasmid by conjugative transformation with a Ralstonia strain, in contrast to the aniline capacity. Additional data suggest that oxidative deamination of chlorinated and nonchlorinated aromatics may be completed by a different set of genes (Boon et al., 2001).

Ecology Comamonas strains have been recovered from sites heavily contaminated with complex organic compounds and heavy metals, and resistance to heavy metals such as cadmium and nickel has been reported (Stoppel and Schlegel, 1995; Kanazawa and Mori, 1996). A strain tentatively identified as C. testosteroni was among the strongest chromate-reducing bacteria found in the cooling water of an electricity plant and may therefore be implicated in the blockage of pipes through precipitation of chromium (III) oxide (Cooke et al., 1995). Most of the isolates from a polychlorinated biphenyl (PCB)-polluted site were Comamonas testosteroni strains capable of degrading biphenyl and a variety of polychlorinated biphenyls (Joshi and Walia, 1995). Many Comamonas strains from various environments are capable of degrading poly-3hydroxybutyrate (PHB; Mergaert and Swings, 1996), and the extracellular PHB depolymerase of one strain has been purified and characterized (Jendrossek et al., 1993). The broad substrate specificity of its 3-αhydroxysteroid dehydrogenases and its occurrence in the intestinal tract of vertebrates have led to suggestions that C. testosteroni and related bacterial strains may contribute to the bioactivation or inactivation of hormones, bile acids and xenobiotics (Oppermann and Maser, 1996a). Comamonas testosteroni possesses steroidinducible hydroxysteroid dehydrogenases and carbonyl reductases which may provide protection against natural and synthetic toxic compounds in soil and the intestinal tract of mammals (Oppermann et al., 1996b).

Pathogenicity Although many Comamonas strains have been isolated from clinical samples and from the hospital environment (Willems et al., 1991), they are regarded as very occasional opportunistic pathogens (Gilardi, 1985), and no evidence of

732

A. Willems and P. De Vos

pathogenic effect on healthy people has been reported.

Applications Because they can break down a wide variety of complex organic compounds (including carboxylic and dicarboxylic aliphatic or unsaturated carboxylic acids, aromatic compounds, and sterols), Comamonas strains contribute to bioremediation processes. Comamonas testosteroni has been used for the degradation of 4-toluenesulfonic acid in a continuously operated fixed-bed biofilm reactor (Khlebnikov and Peringer, 1996). Analysis of the bacterial communities in activated sludge revealed the presence of Comamonas strains (Kaempfer et al., 1996). Comamonas terrigena strain N3H, immobilized in an alginate gel, has been used for the degradation of the anion-active surfactant dihexyl-sulfosuccinate (Huska et al., 1996; Toth et al., 1996; Proksova et al., 1999). Comamonas strains JS46 and JS 47 have been studied in mixed and non-mixed culture immobilized cell reactors for the degradation of m- and p-nitrobenzoate (Goodall et al., 1998b). The experimental data were used for process modeling (Goodall and Peretti, 1998a). A quinohemoprotein ethanol dehydrogenase, which catalyzes the NAD-independent oxidation of a broad range of alcohols to their aldehydes and on to the carboxylic acid, has been purified from ethanol-grown C. testosteroni cells. The enzyme was studied for use in the production of (S)-solketal (2,2-dimethyl-1,3-dioxolane-4methanol) by enantioselective oxidation of racemic solketal (Geerlof et al., 1994; Machado et al., 1999). The active holoenzyme contains pyrroquinoline-quinone, calcium ions, and heme c (De Jong et al., 1995). The encoding gene (qhedh) was cloned, sequenced and expressed in E. coli (Stoorvogel et al., 1996). The enzyme has been applied in electrodes through immobilization in a redox polymer network (Stigter et al., 1997) and can be used for the enantioselective oxidation of secondary alcohols (Godde et al., 1999). The capacity of C. testosteroni BS1310 (pBS1010) to degrade sulfoaromatic compounds resulted in the design of a Clark-type electrode to measure the p-toluene concentration (Makarenko et al., 1999). The ability of C. testosteroni to selectively degrade L-lysine from racemized lysine crystals has been applied in the large-scale production of D-lysine (Takahashi et al., 1997). Finally, a number of studies point to other potential applications of Comamonas strains. Comamonas testosteroni strain CMI 2848 had the highest activity for hydroxylating 3cyanopyridine to 3-cyano-6-hydroxypyridine

CHAPTER 3.2.7

among 4600 isolates screened in a study contributing to the synthesis of new agricultural chemicals (Yasuda et al., 1995). Because hydroxylated N-hetero-aromatic compounds are difficult to produce, the characteristic of a quinolineoxidoreductase of C. testosteroni 63 to mediate the hydroxylation of N-hetero-aromatic compounds such as pyridine to hydroxy-substituted pyrimidines may be of pharmaceutical importance (Fetzner, 1999).

Literature Cited Abalain, J. H., S. Di Stefano, M. L. Abalain-Colloc, and H. H. Floch. 1995. Cloning, sequencing and expression of Pseudomonas testosteroni gene encoding 3alpha-hydroxysteroid dehydrogenase. J. Steroid Biochem. Molec. Biol. 55:233–238. Ahmad, D., J. Fraser, M. Sylvestre, A. Larose, A. Kahn, J. Bergeron, J. M. Juteau, and M. Sondossi. 1995. Sequence of the bphD gene encoding 2hydroxy-6-oxo-(phenyl/ chlorophenyl)hexa-2,4 dienoic acid (HOP/cPDA) hydrolase involved in the biphenyl/polychlorinated biphenyl degradation pathway in Comamonas testosteroni: Evidence suggesting involvement of Ser112 in catalytic activity. Gene 156:69–74. Amann, R., W. Ludwig, R. Schulze, S. Spring, E. Moore, and K.-H. Schleifer. 1996. rRNA-targeted oligonucleotide probes for the identification of genuine and former pseudomonads. Syst. Appl. Microbiol. 19:501–509. Arai, H., Akahira, S., T. Ohishi, and T. Kudo. 1999a. Adaptation of Comamonas testosteroni TA441 to utilization of phenol by spontaneous mutation of the gene for a trans-acting factor. Molec. Microbiol. 33:1132–1140. Arai, H., T. Yamamoto, T. Ohishi, T. Shimizu, T. Nakata, and T. Kudo. 1999b. Genetic organization and characteristics of the 3-(3-hydroxyphenyl)propionic acid degradation pathway of Comamonas testosteroni TA441. Microbiology 145:2813–2820. Arai, H., T. Ohishi, M. Y Chang, and T. Kudo. 2000. Arrangment and regulation of the genes for the meta-pathway enzymes required for the degradation of phenol in Comamonas testosteroni TA441. Microbiology 146: 1707–1715. Baker, M. E. 1996. Bacterial 3-alpha-hydroxysteroid dehydrogenase is homologous to a fusion of bacterial ribosomal L10 and L7/12 genes. J. Steroid Biochem. Molec. Biol. 59:365–366. Ben-Tovim, T., E. Eylan, A. Romano, and R. Stein. 1974. Gram-negative bacteria isolated from external eye infections. Infection 2:162–165. Bergeron, J., D. Ahmad, D. Barriault, A. Larose, M. Sylvestre, and J. Powlowski. 1994. Identification and mapping of the gene translation products involved in the first steps of the Comamonas testosteroni B-356 biphenyl/ chlorobiphenyl degradation pathway. Can. J. Microbiol. 40:743–753. Boon, N., J. Goris, P. De Vos, W. Verstraete, and E. Top. 2000. Bioaugmentation of activated sludge by an indigenous 3-chloroaniline-degrading Comamonas testosteroni strain, I2gfp. Appl. Environ. Microbiol. 66:2906– 2913. Boon, N., J. Goris, P. De Vos, W. Verstraete, and E. Top. 2001. Genetic diversity among 3-chloroaniline- and aniline

CHAPTER 3.2.7 degrading strains of the Comamonadaceae. Appl. Environ. Microbiol. 67:1107–1115. Brothers, P. N., G. Blotny, L. Qi, and R. M. Pollack. 1995. An active site phenylalanine of 3-oxo-d5-steroid isomerase is catalytically important for proton transfer. Biochemistry 34:15453–15458. Byng, G. S., J. L. Johnson, R. J. Whitaker, R. L. Gherna, and R. A. Jensen. 1983. The evolutionary pattern of aromatic amino acid biosynthesis and the emerging phylogeny of pseudomonad bacteria. J. Molec. Evol. 19:272–282. Cabrera, J. E., G. Panzetta-Dutari, J. L. Pruneda, and S. Genti-Raimondi. 1997. A new Comamonas testosteroni steroid-inducible gene: cloning and sequence analysis. J. Steroid Biochem. Molec. Biol. 63:91–98. Chebrou, H., Y. Hurtubise, D. Barriault, and M. Sylvestre. 1999. Heterologous expression and characterization of the purified oxygenase component of Rhodococcus globerulus P6 biphenyl dioxygenase and of chimeras derived from it. J. Bacteriol. 181:4805–4811. Cooke, V. M., M. N. Hughes, and R. K. Poole. 1995. Reduction of chromate by bacteria isolated from the cooling water of an electricity generating station. J. Indust. Microbiol. 14:323–328. Coote, J. G., and H. Hassall. 1973. The degradation of Lhistidine, imidazolyl-L-lactate, and imidazolylpropionate by Pseudomonas testosteroni. Biochem. J. 132:409–422. Dagley, S., and M. D. Patel. 1957. Oxidation of p-cresol and related compounds by a Pseudomonas. Biochem. J. 66:227–233. Davis, G. H. G., and R. W. A. Park. 1962. A taxonomic study of certain bacteria currently classified as Vibrio species. J. Gen. Microbiol. 27:101–119. De Jong, G. A. H., A. Geerlof, J. Stoorvogel, J. A. Jongejan, S. De Vries, and J. A. Duine. 1995. Quinohaemoprotein ethanol dehydrogenase from Comamonas testosteroni: Purification, characterization and reconstitution of the apoenzyme with pyrroloquinoline quinone analogues. Eur. J. Biochem. 230:899–905. De Rore, H., K. Demolder, K. De Wilde, E. Top, F. Houwen, and W. Verstraete. 1994. Transfer of the catabolic plasmid RP4::Tn4371 to indigenous soil bacteria and its effect on respiration and biphenyl breakdown. FEMS Microbiol. Ecol. 15:71–77. De Vos, P., K. Kersters, E. Falsen, B. Pot, M. Gillis, P. Segers, and J. De Ley. 1985. Comamonas Davis and Park 1962 gen. nov., nom. Rev. emend., and Comamonas terrigena Hugh 1962 sp. nov., nom. rev. Int. J. Syst. Bacteriol. 35:443–453. Etchebehere, C., M. I. Errazquin, P. Dabert, R. Moletta, and L. Muxí. 2001. Comamonas nitrativorans sp. nov., a novel denitrifier isolated from a denitrifying reactor treating landfill leachate. Int. J. Syst. Evol. Microbiol. 51:977–983. Fetzner, S. 1999. Bioconversion of pyrimidine by resting cells of quinoline-degrading bacteria. FEMS Microbiol. Lett. 176:291–299. Florin, C., T. Koehler, M. Grandguillot, and P. Plesiat. 1996. Comamonas testosteroni 3-ketosteroid-δ4(5-α)dehydrogenase gene and protein characterization. J. Bacteriol. 178:3322–3330. Galarneault, T. P., and E. Leifson. 1956. Taxonomy of Lophomonas N. Gen. Can. J. Microbiol. 2:102–110. García-Valdés, E., E. Cozar, R. Rotger, J. Lalucat, and J. Ursing. 1988. New naphthalene-degrading marine Pseudomonas strains. Appl. Environ. Microbiol. 54:2478–2485.

Comamonas

733

Geerlof, A., J. Stoorvogel, J. A. Jongejan, E. J. T. M. Leenen, T. J. G. M. Van Dooren, W. J. J. Van Den Tweel, and J. A. Duine. 1994. Studies on the production of (S)-(+)solketal (2,2-dimethyl-1,3-dioxolane-4-methanol) by enantioselective oxidation of racemic solketal with Comamonas testosteroni. Appl. Microbiol. Biotechnol. 42:8–15. Gilardi, G. L. 1971. Characterization of nonfermentative, nonfastidious Gram-negative bacteria encountered in medical bacteriology. J. Appl. Bacteriol. 34:623–644. Gilardi, G. L. 1985. Pseudomonas. In: E. H. Lennette, A. Balows, W. J. Hausler Jr., and H. J. Shadomy (Eds.) Manual of Clinical Microbiology, 4th ed. American Society for Microbiology. Washington, DC. 350–372. Godde, C., M. Liebergesell, and A. Steinbuchel. 1999. Isolation of poly(beta-L-malic acid)-degrading bacteria and purification and characterization of the PNA hydrolase from Comamonas acidovorans strain 7789. FEMS Microbiol. Lett. 173:365–372. Goodall, J. L., and S. W. Peretti. 1998a. Dynamic modeling of meta- and para-nitrobenzoate metabolism by a mixed co-immobilized culture of Comamonas spp. JS46 and JS47. Biotech. Bioeng. 59:507–516. Goodall, J. L., S. M. Thomas, J. C. Spain, and S. W. Peretti. 1998b. Operation of mixed-culture immobilized cell reactors for the metabolism of meta- and paranitrobenzoate by Comamonas sp. JS46 and Comamonas sp. JS47. Biotechnol. Bioeng. 59:21–27. Goyal, A. K., and G. J. Zylstra. 1996. Molecular cloning of novel genes for polycyclic aromatic hydrocarbon degradation from Comamonas testosteroni GZ29. Appl. Environ. Microbiol. 62:230–236. Gray, P. H. H., and H. G. Thornton. 1928. Soil bacteria that decompose certain aromatic compounds. Zentralbl. Bakteriol. Parasitenk. Abt. 2 73:74–96. Grones, J., and K. Turna. 1995. Transformation of microorganisms with the plasmid vector with the replicon from pAC11 from Acetobacter pasteurianus. Biochem. Biophys. Res. Comm. 206:942–947. Gumaelius, L., G. Magnusson, B. Petterson, and G. Dalhammar. 2001. Comamonas denitrificans sp.nov., an efficient denitrifying bacterium isolated from activated sludge. Int. J. Syst. Evol. Microbiol. 51:999–1006. Gunther, K. A. 1894. Über einen neuen in Erdboden gefundenen Kommabacillus. Zentralbl. Bakteriol. Parasitenkd. 16:746–747. He, Z. Q., and J. C. Spain. 1999. Comparison of the downstream pathways for degradation of nitrobenzene by Pseudomonas pseudoalcaligenes JS45 (2-aminophenol pathway) and by Comamonas sp. JS765 (catechol pathway). Arch. Microbiol. 171:309–316. Hein, P., J. Powlowski, D. Barriault, Y. Hurtubise, D. Ahamad, and M. Sylvestre. 1998. Biphenyl-associated meta-cleavage dioxygenase from Comamonas testosteroni B-356. Can. J. Microbiol. 44:42–49. Hollender, J., W. Hopp, and W. Dott. 1997. Degradation of 4-chlorophenol via the meta cleavage pathway by Comamonas testosteroni JH5. Appl. Environ. Microbiol. 63:4567–4572. Hrywna, Y., T. V. Tsoi, O. V. Maltseva, J. F. Quensen, and J. M. Tiedje. 1999. Construction and characterization of two recombinant bacteria that grow on ortho- and parasubstituted biphenyls. Appl. Environ. Microbiol. 65:2163–2169.

734

A. Willems and P. De Vos

Hugh, R. 1962. Comamonas terrigena comb. nov. with proposal of a neotype an request for an opinion. Int. Bull. Bacteriol. Nomencl. Tax. 12:33–35. Hugh, R. 1965. A comparison of Pseudomonas testosteroni and Comamonas terrigena. Int. Bull. Bacteriol. Nomencl. Tax. 15:125–132. Hungerer, C., B. Troup, U. Roemling, and D. Jahn. 1995. Regulation of the hemA gene during 5-aminolevulinic acid formation in Pseudomonas aeruginosa. J. Bacteriol. 177:1435–1443. Hurtubise, Y., D. Barriault, J. Powlowski, and M. Sylvestre. 1995. Purification and characterization of the Comamonas testosteroni B-356 biphenyl dioxygenase components. J. Bacteriol. 177:6610–6618. Hurtubise, Y., D. Barriault, and M. Sylvestre. 1996. Characterization of active recombinant His-tagged oxygenase component of Comamonas testosteroni B-356 biphenyl dioxygenase. J. Biol. Chem. 271:8152–8156. Huska, J., I. Zavdska, D. Toth, M. Dobrotova, and P. Gemeiner. 1996. Immobilization of surfactant degrading bacteria in alginate gel. Biologia (Bratislava) 51:279– 283. Jayarao, B. M., and L. Wang. 1999. A study on the prevalence of gram-negative bateria in bulk tank milk. J. Dairy Sci. 82:2620–2624. Jendrossek, D., I. Knoke, R. B. Habibian, A. Steinbüchel, and H. G. Schlegel. 1993. Degradation of poly(3-hydroxybutyrate), PHB, by bacteria and purification of a novel PHB depolymerase from Comamonas sp. J. Environ. Polymer Degrad. 1:53–63. Jendrossek, D., M. Backhaus, and M. Andermann. 1995. Characterization of the extracellular poly(3-hydroxybutyrate) depolymerase of Comamonas sp. and of its structural gene. Can. J. Microbiol. 41:160–169. Joshi, B., and S. Walia. 1995. Characterization by arbitrary primer polymerase chain reaction of polychlorinated biphenyl (PCB)-degrading strains of Comamonas testosteroni isolated from PCB-contaminated soil. Can. J. Microbiol. 41:612–619. Junker, F., E. Saller, H. R. S. Oppenberg, P. M. H. Kroneck, T. Leisinger, and A. M. Cook. 1996. Degradative pathways for p-toluenecarboxylate and p-toluenesulfonate and their multicomponent oxygenases in Comamonas testosteroni strains PSB-4 and T-2. Microbiology 142:2419–2427. Junker, F., and A. M. Cook. 1997a. Conjugative plasmids and the degradation of arylsulfonates in Comamonas testosteroni. Appl. Environ. Microbiol. 63:2403–2410. Junker, F., R. Kiewitz, and A. M. Cook. 1997b. Characterization of the p-toluenesulfonate operon tsaMBCD and tsaR in Comamonas testosteroni T-2. J. Bacteriol. 179:919–927. Kaempfer, P., R. Erhart, C. Beimfohr, J. Boehringer, M. Wagner, and R. Amann. 1996. Characterization of bacterial communities from activated sludge: Culture-dependent numerical identification versus in situ identification using group- and genus-specific rRNA-targeted oligonucleotide probes. Microb. Ecol. 32:111–121. Kanazawa, S., and M. Mori. 1996. Isolation of cadmiumresistant bacteria and their resistance mechanisms. Part I: Isolation of CD-resistant bacteria from soils contaminated with heavy metals. Soil Sci. Plant Nutr. 42:725– 730. Kertesz, M. A. 2000. Riding the sulfur cycle—metabolism of sulfonates and sulfate esters in Gram-negative bacteria. FEMS Microbiol. Rev. 24:135–175.

CHAPTER 3.2.7 Khlebnikov, A., and P. Peringer. 1996. Biodegradation of ptoluenesulphonic acid by Comamonas testosteroni in an aerobic counter-current structured packing biofilm reactor. Water Sci. Technol. 34:257–266. Krieg, N. R., and J. G. Holt (Eds.). 1984. Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. Laitinen, S., M. Linnainmaa, J. Laitinen, H. Kiviranta, M. Reiman, and J. Liesivuori. 1999. Endotoxins and IgG antibodies as indicators of occupational exposure to the microbial contaminants of metal-working fluids. Int. Arch. Occup. Environ. Health. 72:443–450. Lee, J.-H., T. Omori, and T. Kodama. 1994. Identification of the metabolic intermediates of phthalate by Tn5 mutants of Pseudomonas testosteroni and analysis of the 4,5-dihydroxyphthalate decarboxlase gene. J. Ferment. Bioeng. 77:583–590. Lehmann, K. B., and R. Neumann. 1927. Bakteriologie insbesondere bakteriologische Diagnostiek. II. Allgemeine und spezielle Bakteriologie, 7th ed. J. F. Lehmann. Munich, Germany. 548. Lemmer, H., A. Zaglauer, A. Neef, H. Meier, and R. Amann. 1997. Denitrification in a methanol-fed fixedbed reactor. Part 2: Composition and ecology of the bacterial community in the biofilms. Water Res. 31:1903–1908. Levy-Schil, S., F. Soubrier, A.-M. Crutz-Le Coq, D. Faucher, J. Crouzet, and D. Petre. 1995. Aliphatic nitrilase from a soil isolated Comamonas testosteroni sp.: Gene cloning and overexpression, purification and primary structure. Gene 161:15–20. Locher, H. H., Poolman, B., Cook, A. M., and Konings, W. N. 1993. Uptake of 4-toluene sulfonate by Comamonas testosteroni T-2. J. Bacteriol. 175:1075–1080. Machado, S. S., A. Jongejan, A. Geerlof, J. A. Jongejan, and J. A. Duine. 1999. Entropic and enthalpic contributions to the enantioselectivity of quinohaemoprotein alcohol dehydrogenases from Acetobacter pasteurianus and Comamonas testosteroni in the oxidation of primary and secondary alcohols. Biocatal. Biotrans. 17:179–207. Makarenko, A. A., S. V. Balashov, T. N. Kuvichkina, P. V. Il’yasov, and A. N. Reshetilov. 1999. A biosensor of sulfoaromatic compounds based on the strain Comamonas testosteroni BS1310 (pBS101). Appl. Biochem. Microbiol. 35:375–379. Mergaert, J., and J. Swings. 1996. Biodiversity of microorganisms that degrade bacterial and synthetic polyesters. J. Indust. Microbiol. 17:463–469. Meyer, O., E. Stackebrandt, and G. Auling. 1993. Reclassification of ubiquinone Q-10 containing carboxidotrophic bacteria: transfer of “[Pseudomonas] carboxydovorans” OM5T to Oligotropha, gen. nov., as Oligotropha carboxydovorans, comb. nov., transfer of “[Alcaligenes] carboxydus” DSM 1086T to Carbophilus, gen. nov., as Carbophilus carboxidus, comb. nov., transfer of “[Pseudomonas] compransoris” DSM 1231T to Zavarzinia, gen. nov., as Zavarzinia compransoris, comb. nov., and amended descriptions of the new genera. Syst. Appl. Microbiol. 16:390–395. Moebus, E., M. Jahn, R. Schmid, D. Jahn, and E. Maser. 1997. Testosterone-regulated expression of enzymes involved in steroid and aromatic hydrocarbon catabolism in Comamonas testosteroni. J. Bacteriol. 179:5951–5955. Morgan, C. A., and R. C. Wyndham. 1996. Isolation and characterization of resin acid degrading bacteria found

CHAPTER 3.2.7 in effluent from a bleached kraft pulp mill. Can. J. Microbiol. 42:423–430. Mudd, S., and S. Warren. 1923. A readily cultivable vibrio, filterable through Berkefeld “V” candles, Vibrio percolans (new species). J. Bacteriol. 8:447–458. Nozhevnikova, A. N., and G. A. Zavarzin. 1974. On the taxonomy of CO-oxidizing Gram-negative bacteria. Izv. Akad. Nauk SSSR, Ser. Biolog. 3:436–440. Oppenberg, H. R. S., G. Chen, T. Leisinger, and A. M. Cook. 1995. Regulation of the degradative pathways from 4toluenesulphonate and 4-toluenecarboxylate to protocatechuate in Comamonas testosteroni T-2. Microbiology 141:1891–1899. Oppermann, C. T., and E. Maser. 1996a. Characterization of a 3-α-hydroxysteroid dehydrogenase/carbonyl reductase from the Gram-negative bacterium Comamonas testosteroni. Eur. J. Biochem. 241:744–749. Oppermann, U. C. T., I. Belai, and E. Maser. 1996b. Antibiotic resistance and enhanced insecticide catabolism as consequence of steroid induction in the Gram-negative bacterium Comamonas testosteroni. J. Steroid Biochem. Molec. Biol. 58:217–223. Oppermann, U. C. T., C. Filling, K. D. Berndt, B, Persson, J. Benach, R. Ladenstein, and H. Joernvall. 1997. Active site directed mutagenesis of 3β/17β-hydroxysteroid dehydrogenase establishes differential effects on shortchain dehydrogenase/reductase reactions. Biochemistry (Wash.) 36:34–40. Palleroni, N. J. 1984. Genus I. Pseudomonas Migula 1894. In: N. R. Krieg and J. G. Holt (ed.) Bergey’s manual of systematic bacteriology. Williams and Wilkins. Baltimore, MD. 1:141–199. Parales, R. E., T. A. Ontl, and D. T. Gibson. 1997. Cloning and sequence analysis of a catechol 2,3-dioxygenase gene from the nitrobenzene-degrading strain Comamonas sp. JS765. J. Indust. Microbiol. Biotechnol. 19:385– 391. Patureau, D., N. Bernet, and R. Moletta. 1997. Combined nitrification and denitrification in a single aerated reactor using the aerobic denitrifier Comamonas sp. strain SGLY2. Water Res. 31:1363–1370. Petruschky, J. 1896. Bacillus faecalis alcaligenes (n. sp.). Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. I 19:187–191. Pickett, M. J., and J. R. Greenwood. 1986. Pseudomonas alcaligenes and Pseudomonas testosteroni: characterization and identification. Curr. Microbiol. 13:197–201. Proksova, M., D. Sladekova, B. Uhnakova, J. Augustin, and D. Toth. 1999. Primary biodegradtion and assimilation of dialkyl sulphosuccinate by Comamonas terrigena N3H. Biologia 54:671–677. Quinteros, R., S. Goodwin, R. W. Lenz, and W. H. Park. 1999. Extacellular degradation of medium chain length poly(beta-hydroxyalkanoates) by Comamonas sp. Int. J. Biol. Macromol. 25:135–143. Rubin, S. J., P. A. Granato, and B. L. Wasilauskas. 1985. Glucose-nonfermenting Gram-negative bacteria. In: Lennette, Balows, Hausler and Shadomy (Eds.) Manual of clinical microbiology, 4th Ed. American Society for Microbiology. Washington, DC. 330–349. Schach, S., B. Tshisuaka, S. Fetzner, and F. Lingens. 1995. Quinoline 2-oxidoreductase and 2-oxo-1,2-dihydroquinoline 5,6-dioxygenase from Comamonas testosteroni 63: the first two enzymes in quinoline and 3-methylquinoline degradation. Eur. J. Biochem. 232: 536–544.

Comamonas

735

Schlaefi, H. R., M. A. Weiss, T. Leisinger, and A. M. Cook. 1994. Terephthalate 1,2-dioxygenase system from Comamonas testosteroni T-2: Purification and some properties of the oxygenase component. J. Bacteriol. 176:6644– 6652. Shinomiya, M., T. Iwata, K. Kasuya, and Y. Doi. 1997. Cloning of the gene for poly(3-hydroxybutyric acid) depolymerase of Comamonas testosteroni and functional analysis of its substrate-binding domain. FEMS Microbiol. Lett. 154:89–94. Skerman, V. B. D., V. McGowan, and P. H. A. Sneath. 1980. Approved lists of bacterial names. Int. J. Syst. Bacteriol. 30:225–420. Springael, D., J. van Thor, H. Goorissen, A. Ryngaert, R. De Baere, P. Van Hauwe, L. C. M. Commandeur, J. R. Parsons, R. De Wachter, and M. Mergeay. 1996. RP4::Mu3A-mediated in vivo cloning and transfer of a chlorobiphenyl catabolic pathway. Microbiology 142: 3283–3293. Stanier, R. Y., N. J. Palleroni, and M. Doudoroff. 1966. The aerobic pseudomonads: A taxonomic study. J. Gen. Microbiol. 43:159–271. Stigter, E. C. A., G. A. H. De Jong, J. A. Jongejan, J. A. Duine, J. P. Van Der Lugt, and W. A. C. Somers. 1997. Electron transfer and stability of a quinohaemoprotein alcohol dehydrogenase electrode. J. Chem. Technol. Biotechnol. 68:110–116. Stoorvogel, J., D. E. Kraayveld, C. A. Van Sluis, J. A. Jongejan, S. De Vries, and J. A. Duine. 1996. Characterization of the gene encoding quinohaemoprotein ethanol dehydrogenase of Comamonas testosteroni. Eur. J. Biochem. 235:690–698. Stoppel, R. D., and H. G. Schlegel. 1995. Nickel-resistant bacteria from anthropogenically nickel-polluted and naturally nickel-percolated ecosystems. Appl. Environ. Microbiol. 61:2276–2285. Sylvestre, M., Y. Hurtubise, D. Barriault, J. Bergeron, and D. Ahmad. 1996a. Characterization of active recombinant 2,3-dihydro-2,3-dihydroxybiphenyl dehydrogenase from Comamonas testosteroni B-356 and sequence of the encoding gene (bphB). Appl. Environ. Microbiol. 62:2710–2715. Sylvestre, M., M. Sirois, Y. Hurtubise, J. Bergeron, D. Ahmad, F. Shareck, D. Barriault, I. Guillemette, and J. M. Juteau. 1996b. Sequencing of Comamonas testosteroni strain B356 biphenyl/chlorobiphenyl deoxygenase genes: Evolutionary relationships among Gram-negative bacterial biphenyl dioxygenases. Gene 174:195–202. Takahashi, E., M. Furui, and T. Shibatani. 1997. Scale-up of D-lysine production from L-lysine by successive chemical racemization and microbial asymmetric degradation. Biotechnol. Lett. 19:245–249. Talalay, P., M. Mollomo Dobson, and D. F. Tapley. 1952. Oxidative degradation of testosterone by adaptive enzymes. Nature (London) 170:620–621. Tamaoka, J., D.-M. Ha, and K. Komagata. 1987. Reclassification of Pseudomonas acidovorans den Dooren de Jong 1926 and Pseudomonas testosteroni Marcus and Talalay 1956 as Comamonas acidovorans comb. nov. and Comamonas testosteroni comb. nov., with an emended description of the genus Comamonas. Int. J. Syst. Bacteriol. 37:52–59. Tazuke, Y., K. Matsuda, K. Adachi, and Y. Tsukada. 1994. Purification and properties of bile acid sulfate sulfatase from Pseudomonas testosteroni. Biosci. Biotechnol. Biochem. 58:889–894.

736

A. Willems and P. De Vos

Teramoto, M., H. Futamata, S. Harayama, and K. Watanabe. 1999. Characterization of a high-affinity phenol hydroxylase from Comamonas testosteroni R5 by gene cloning, and expression in Pseudomonas aeruginosa PAO1c. Molec. Gen. Genet. 262:552–558. Toth, D., J. Huska, I. Zavadska, and M. Dobrotova. 1996. Effect of bacterial starvation on surfactant biotransformation. Folia Microbiol. 41:477–479. Vanbrabant, J., P. De Vos, M. Vancanneyt, J. Liessens, W. Verstraete, and K. Kersters. 1993. Isolation and identification of autotrophic and heterotrophic bacteria from an autohydrogenotrophic pilot-plant for denitrification of drinking-water. Syst. Appl. Microbiol. 16:471–482. Vaneechoutte, M., R. Rossau, P. De Vos, M. Gillis, D. Janssens, N. Paepe, A. De Rouck, T. Fiers, G. Claeys, and K. Kersters. 1992. Rapid identification of bacteria of the Comamonadaceae with amplified ribosomal DNArestriction analysis (ARDRA). FEMS Microbiol. Lett. 93:227–234. Wen, A, M. Fegan, C. Hayward, and S. Chakraborty, and L. I. Sly. 1999. Phylogenetic relationships among members of the Comamonadaceae, and description of Delftia acidovorans (den Dooren de Jong 1926, and Tamaoka et al. 1987) gen. nov., comb. nov. Int. J. Syst. Bacteriol. 49:567–576. Willems, A., B. Pot, E. Falsen, P. Vandamme, M. Gillis, K. Kersters, and J. De Ley. 1991. Polyphasic taxonomic study of the emended genus Comamonas: Relationship

CHAPTER 3.2.7 to Aquaspirillum aquaticum, E. Falsen group 10, and other clinical isolates. Int. J. Syst. Bacteriol. 41:427–444. Xiong, G. M., H. J. Martin, C. Schafers, and E. Maser. 2001. A model on the regulation of 3 alpha-hydroxysteroid dehydrogenase/carbonyl reductase expression in Comamonas testosteroni. Chemico-biol. Interact. 13:723–736. Yap, L. F., Y. K. Lee, and C. L. Poh. 1999. Mechanism for phenol tolerance in phenol-degrading Comamonas testosteroni strains. Appl. Microbiol. Biotechnol. 51:833– 840. Yasuda, M., T. Sakamoto, R. Sashida, M. Ueda, Y. Morimoto, and T. Nagasawa. 1995. Microbial hydroxylation of 3cyanopyridine to 3-cyano-6-hydroxypyridine. Biosci. Biotechnol. Biochem. 59:572–575. Zhao, Q., Abeygunawardana, C., Talalay, P., and Mildvan, A. S. 1996. NMR evidence for the participation of a lowbarrier hydrogen bond in the mechanism of d5-3ketosteroid isomerase. Proc. Natl. Acad. Sci. 93:8220– 8224. Zhao, Q., Abeygunawardana, C., and Mildvan, A. S. 1997. NMR studies of secondary structure in isolation and the steroid binding site of d5-3-ketosteroid isomerase in complexes with diamagnetic and paramagnetic steroids. Biochemistry (Wash.) 36:3458–3472. Zhao, J. S., and O. P. Ward. 1999. Microbial degradation of nitrobenzene and mono-nitrophenol by bacteria enriched from municipal activated sludge. Can. J. Microbiol. 45:427–432.

Prokaryotes (2006) 5:737–746 DOI: 10.1007/0-387-30745-1_32

CHAPTER 3.2.8 ehT

areneG

mu i re t cabomorhC

dna

mu i re t cabon i htna J

The Genera Chromobacterium and Janthinobacterium MONIQUE GILLIS AND JOZEF DE LEY

Purple-pigmented bacteria have been described since the end of the 19th century; they were reported as discoloring a variety of natural materials and occasionally as the causative agent of septicemia in humans and animals. Bacteria producing purple and violet colonies due to the production of a nondiffusible pigment, violacein, were classified in a redefined genus Chromobacterium by Buchanan (1918). The structure of violacein, an indole derivative produced via the oxidation of tryptophan, is shown in Fig. 1. It can be easily identified (Johnson and Beer, 1971) spectrophotometrically from the following properties: 1) in ethanolic solution it has an absorption maximum at 579 nm and a minimum at 430 nm; 2) by adding 10% (v/v) H2SO4 the solution turns green with an absorption maximum at 700 nm; and 3) when NaOH is added to an ethanolic solution, the solution turns green and afterwards reddish brown. Recently, violacein has also been characterized by nuclear magnetic resonance spectroscopy and mass spectrometry (Riveros et al., 1988). The pigment is only abundantly produced when tryptophan is available in the culture medium. Although violacein is produced by only a few groups of bacteria, its presence does not necessarily indicate a close relationship between these organisms. An extensive phenotypic study (Sneath, 1956, 1960, 1974) within Chromobacterium, as defined by Buchanan (1918), provided evidence of two groups within this genus: a fermentative and mesophilic one (growth at 37°C but not at 4°C) and a nonfermentative and psychrophilic one (growth at 4°C but not at 37°C), for which he proposed two species, Chromobacterium violaceum and [Chromobacterium] lividum, respectively. However, these two species have been reported as being not closely related (Moffet and Colwell, 1968; Sneath, 1974) and since the latter was misnamed, in this chapter its epithet and all other misnamed epithets shall be enclosed in square brackets. Violacein is not a good taxonomic marker as shown by the fact that not only have nonpigmented Chromobacterium violaceum and [Chromobacterium] lividum strains been isolated, but also since marine, violacein-producing strains were isolated, which were assigned on phenotypic grounds to the genus Alteromonas (Gauthier, 1976, 1982). Later, Van Landschoot and De Ley (1983) confirmed that these marine strains were genuine Alteromonas. The relationships between the violacein-producing bacteria classified in Chromobacterium (Sneath, 1974) could finally be elucidated by De Ley et al. (1978). DNA-rRNA hybridizations within and between the violaceum and the lividum taxons and with different other Gram-negative genera demonstrated clearly that these purple-pigmented bacteria represent two tight but separate groups within rRNA This chapter was taken unchanged from the second edition.

superfamily III (see also De Ley’s introduction to the Proteobacteria in The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition) which are less related to each other than to other genera from this rRNA superfamily. In consequence, a new genus Janthinobacterium with one species J. lividum was created for the lividum taxon and the genus name Chromobacterium was restricted to the mesophilic, fermentative violaceum taxon (De Ley et al., 1978). Chromobacterium constitutes a separate rRNA branch on which all violaceum strains cluster with a Tm(e) of 76.5 to 80°C. Janthinobacterium constitutes a separate rRNA subbranch which is closely related to the rRNA subbranch containing [Pseudomonas] rubrisubalbicans and a yet unnamed group of bacteria isolated from clinical material (see Fig. 6 in The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition). The Janthinobacterium-[Pseudomonas] rubrisubalbicans rRNA branch is more closely related to the solanacearum rRNA branch than to Chromobacterium, emphasizing the deep phylogenetic gap between Chromobacterium and Janthinobacterium. Cataloging results (Woese et al., 1984) confirm the separate position of both genera in the beta group (Stackebrandt et al., 1988). In this chapter, Chromobacterium and Janthinobacterium are treated together merely on historical grounds. Moss et al. 1978 described a new group of violaceincontaining, fermentative, psychrophilic, fresh water bacteria and assigned them to a new species fluviatile in the genus Chromobacterium. It differs from C. violaceum by a thin flat, spreading colony type, some other phenotypic features (see below) and % GC values of 50 to 52. Within C. violaceum the GC content varies from 65 to 68 mol%, indicating at once that these new organisms cannot be members of Chromobacterium. DNA-rRNA hybridizations (Moss and Bryant, 1982) demonstrated that [Chromobacterium]* fluviatile is not closely related to the C. violaceum rRNA cluster. With a Tm(e) of 70 to 71°C vs. the C. violaceum rRNA probe, they occupy a separate position and are further removed from C. violaceum than the latter species is from the Neisseriaceae (Rossau et al., 1989). Comparison of their 16S rRNA sequences corroborates this conclusion (Dewhirst et al., 1989). Therefore, these organisms cannot belong in Chromobacterium; they have thus been misnamed and shall not be discussed here further. The differentiation of these three groups of violacein-producing organisms is given in Table 1.

*Since this manuscript was submitted, Logan (1989) proposed a monospecific genus Iodobacter to accomodate the [Chromobacterium] fluviatile group. Their phenotypic results on Iodobacter and the “atypical, Janthinobacterium lividum strains” are integrated in Table 1.

738

M. Gillis and J. de Ley

CHAPTER 3.2.8

Chromobacterium The following description of the genus is based on that of Chromobacterium violaceum by Sneath (1974, 1984). The genus Chromobacterium consists of Gram-negative, oxidase-positive, catalase-positive rods with rounded ends which are usually motile by both a single polar flagellum and by one to four lateral flagella. The two types of flagella differ in their wave types, staining capacity, and antigenic characteristics. Chromobacteria are facultative anaerobes and produce violet colonies on solid media; a violet ring is formed in liquid media at the surface with a fragile pellicle. Growth is best at 30 to 35°C; the minimum growth temperature is 10 to 15°C and the maximum is 40°C (20% of the strains can grow at 44°C). They are chemoorganotrophs

using mainly carbohydrates fermentatively. Nitrate and usually nitrite are reduced, mostly with a variable gas production. They grow on ordinary media and are resistant to benzylpenicillin (10 µg/ml) and to vibriostatic agent 0/129 [2,4-diamino-6,7-diisopropylpteridine (30 µg/ disc)]. The GC content of the DNA is 65 to 68 mol%. Other characteristics are as described by Sneath (1984) for C. violaceum. The type and sole species is C. violaceum. Between representative strains of this species, more than 83% DNA binding was found (De Ley et al., 1978). C. violaceum is a saprophyte from soil and water and is normally considered as nonpathogenic for humans. Occasionally, however, it is an opportunistic pathogen of extreme virulence for humans and animals (see below).

Habitats H N

C. violaceum is mainly found in soil and water (Sneath, 1956), where it usually constitutes only a minor component of the total microflora. It is found in temperate but more frequently in tropical and subtropical regions. It was suggested that it can be isolated more frequently from soil than from water (Corpe, 1951; Moss and Ryall, 1981). Its role in the rhizosphere of plants is not clear, although Hussain and Van cˇ ura (1970) isolated C. violaceum strains from rhizosphere soil of maize and demonstrated that the inoculation of maize

O

HO

N H

O

N H

Fig. 1. Structure of violacein.

Table 1. Characteristics differentiating the groups of violacein-producing organisms. Character Growth at 4°C Growth at 37°C Fermentation of glucose (O/F test) Anaerobic growth Cyanide produced Turbid zone on egg yolk agar (lecithinase) Acid from Trehalose L-arabinose D-cellobiose D-galactose Gluconate D-maltose N-acetylglucosamine Lactate utilization Casein hydrolysis Esculin hydrolysis Arginine decarboxylase GC content (mol%) a

Chromobacterium

Janthinobacterium

Iodobacter [Chromobacterium] fluviatilis

− + Fb + + +

+ (96%)a − 0c (90%) − (96%) − −

+ − F + − (+)d

+ − − − + − + + + − + 65–68

− (82%) + + (99%) + (96%) − (99%) + (96%) − (99%) + (99%) − or (+) + (80%) − 65–66

+ − − − + (96%) + + − + − − 50–52

Numbers in parentheses indicate the % of strains giving the indicated reaction. F, fermented. c O, oxidized. d (+) weakly positive. The phenotypic data are from Sneath (1984), Moss et al. (1978) and Logan (1989). The GC values are from De Ley et al. (1978) and from Moss et al. (1978) for Iodobacter [Chromobacterium] fluviatilis. b

CHAPTER 3.2.8

The Genera Chromobacterium and Janthinobacterium

seeds with such C. violaceum strains significantly increased the yield of dry matter of the plant. C. violaceum could not be recovered from food products (Koburger and May, 1982). C. violaceum has been isolated from severe infections in humans and animals; water or soil contaminated with C. violaceum is mostly the causal agent. Although a C. violaceum infection in humans can result in mild diarrhea with a subsequent recovery (Sneath et al., 1953), it is mostly fatal for both humans and animals. A review of the earlier literature concerning chromobacteriosis was given by Sneath (1960). Later rare fatal cases of infection with C. violaceum were reported in different countries, i.e., the United States (Johnson et al., 1971), Vietnam (Ognibene and Thomas, 1970), Cuba (Machin Villafranca et al., 1986), Taiwan (Wu et al., 1986), Argentina (Kaufman et al., 1986), Nigeria (Onile et al., 1984), Brazil (Petrillo et al., 1984), and Australia (Wilkey and McDonald, 1983). The symptoms of the infection can vary considerably but lead mostly to septicemia, causing a rapid death. In the different case reports the symptoms were liver and lung abscesses (Sneath, 1960), meningitis (Shetty et al., 1987), adenitis (Sorensen et al., 1985), periorbital cellulitis (Simo et al., 1984; Feldman et al., 1984), toxemia, and multiple skin abscesses (Petrillo et al., 1984). Only exceptionally nonfatal chromobacterial sepsis has been reported as, i.e., a case of multiple liver abscesses (Suarez et al., 1986) or a systemic chromobacteriosis (Victorica et al., 1974). The disease can be cured by tetracyclines if treated at an early stage (Moss and Ryall, 1981). A recent comparison of the in vitro activity of 25 antimicrobial agents against clinical strains of C. violaceum showed that ciprofloxacin, norfloxacin, and perfloxacin are also highly active (Aldridge et al., 1988). For some eye infections, early recognition and aggressive treatment (with tetracycline, chloramphenicol, and cotrinoxazole) may result in a higher cure rate (Feldman et al., 1984). Infections in animals were also described and studied intensively, i.e., in swine (Laws and Hall, 1964; Liu et al., 1988), Malayan gibbons, a Malayan sun bear (Groves et al., 1969), an Assam macaque (McClure and Chang, 1976), a dog (Gogolewski, 1983), and monkeys (Collins et al., 1985). Virulence for mice and the pathology of infections appear to be similar for both nonpigmented and pigmented C. violaceum strains (Sivendra and Tan, 1977). Virulent C. violaceum strains produce an endotoxin which is more reactive than that from avirulent strains and it is suggested that the virulent strains are protected from phagocytic attack by elevated levels of superoxide dismutase and catalase (Miller et al.,

739

1988). Experimental infections with C. violaceum or with its endotoxin were studied in mice and pigs (Zwang et al., 1987; Liu et al., 1988).

Enrichment and Isolation The medium and conditions used to isolate and enumerate C. violaceum can strongly influence their recovery (Ryall and Moss, 1975; Koburger and May, 1982). Chromobacteria can grow easily on common laboratory media such as nutrient agar or GYCA (1% glucose, 0.5% yeast extract, 3% CaCO3, and 2% agar in distilled water), and violacein is produced in the presence of oxygen. Sneath (1960) concluded that blood agar base was more suitable for the isolation of small numbers of C. violaceum from medical sources, because they can be inhibited by the presence of peroxide which can be present in nutrient agar. For isolation from soil and water, the common media are not so suitable because chromobacteria represent only a minor part of the present microflora, so that large numbers of other bacteria can overgrow or prevent chromobacteria from growing well and producing violacein, which is an important feature in screening. Therefore, an enrichment procedure or a selective medium is recommended. In most cases, the selective media enrich for Chromobacterium as well as for Janthinobacterium and [Chromobacterium] fluviatile. The latter two taxa can be differentiated from C. violaceum afterwards by their phenotypic features (see Table 1). Corpe (1951) described the following simple enrichment technique: “Five grams samples of soil were placed in sterile Petri dishes and soaked with sterile distilled water. Sterile polished or precooked rice grains were sprinkled over the surface of the soil and the plates incubated at 23– 25°C for five days. At the end of the incubation period, the rice grains were partially covered with purple membranous growth indicating the presence of Chromobacterium sp.” Ryall and Moss (1975) used a selective medium based on quarter strength nutrient agar to count chromobacteria and janthinobacteria in water samples: Composition of the Medium of Ryall and Moss (1975). Beef extract (Oxoid) Yeast extract Peptone NaCl Agar Distilled water

0.25 g 0.5 g 1.25 g 1.25 g 20 g 1 liter

To the molten agar medium (45°C), filter-sterilized solutions of colistin and sodium deoxycholate were added to give final concentrations of 15 µg/ml and 0.3 mg/ml, respectively. For counting chromobacteria and janthinobacteria in soil samples, cycloheximide was also added to

740

M. Gillis and J. de Ley

give a final concentration of 30 µg/ml to prevent overgrowth by fungi.

Composition of the Medium of Keeble and Cross (1977). An improved selective medium (Keeble and Cross, 1977) is based on Bennett’s agar: Yeast extract (Difco) Beef extract (Oxoid) Bacto-casitone (Difco) Glucose Agar (Lab. M no. 2) Distilled water

1g 1g 2g 10 g 18 g 1 liter

To the molten agar medium were added filter-sterilized solutions of neomycin hydrochloride, cycloheximide, and nystatin to give a final concentration of 50 µg/ml of each. This medium is described as improving the pigmentation.

After comparing several media, Koburger and May (1982) reported that Bennett’s agar was best for the isolation of both C. violaceum and Janthinobacterium lividum when incubated at 25°C but that at 35°C Aeromonas membrane agar gave the highest recoveries of C. violaceum. Aeromonas membrane agar (Rippey and Cabelli, 1979) is prepared as follows: Tryptose Trehalose Yeast extract NaCl KCl MgSO4 ·7H 2O FeCl3 ·6H 2O Bromothymol blue Deionized water

0.5 g 0.5 g 0.2 g 0.3 g 0.2 g 0.02 g 0.01 g 0.004 g 100 ml

The ingredients are dissolved at room temperature, the pH is adjusted to 8.0 with 10 N NaOH, 1.5 g agar is added, and the mixture is autoclaved at 121°C for 15 min. After the addition of 1.0 ml ethanol, the mixture is cooled to 50°C, and 2 mg ampicillin and 10 mg desoxycholate are added. It is recommended that the poured plates be stored in the dark at 4°C and used within 6 weeks.

For inoculation, 0.1-ml samples are pipetted onto the surface and spread. They are allowed to dry for 30 min and incubated at 25 and 35°C for 7 and 5 days, respectively. All purple colonies developing on the plate were isolated and identified. Koburger and May (1982) suggested that the failure in other studies (Ryall and Moss, 1975; Keeble and Cross, 1977) to recover C. violaceum from soil and water and their own failure to isolate it from a limited number of food products can be related to the temperature sensitivity of these organisms. C. violaceum dies rapidly at 4°C. Chromobacterium can also be isolated on a citrate-ammonium salts medium (Moss and Ryall, 1981) as described for Janthinobacterium further in this chapter. Preservation of Strains The organisms can survive for several years in dilute peptone water (0.1% peptone) at room temperature. On

CHAPTER 3.2.8

GYCA or nutrient agar slants they can be kept for some months. Lyophilization and L-drying can be used to preserve them for longer periods. Freezing in nutrient broth containing 15% glycerol with storing at −80°C or in liquid N2 is also recommended.

Identification When definitely pigmented, Chromobacterium has only to be differentiated from other violacein-producing nonmarine strains belonging in the genus Janthinobacterium or to [Chromobacterium] fluviatile. The differentiating features are given in Table 1. Chromobacterium differs from Alteromonas luteoviolacea in its higher GC content and its ability to use citrate, glucose, fructose, and mannose (Gauthier, 1976, 1982). Nonpigmented C. violaceum strains can be confused with members of Vibrio and Aeromonas. However, C. violaceum differs from Aeromonas in its flagellar morphology, its negative reaction for indole and for methyl red, and its production of HCN. Moreover, C. violaceum differs from Vibrio in its GC content and by a negative response to the lysine and ornithine decarboxylase tests but by a positive reaction in the arginine decarboxylase test. Unlike C. violaceum, most strains of the genera Aeromonas and Vibrio are able to use D-mannitol as a carbon source (Sneath, 1984; Sivendra and Tan, 1977). Singh et al. (1988) used a DNA dot blot technique and a DNA probe from their nonpigmented isolate to differentiate the latter strain from Aeromonas and to identify it as C. violaceum. Members of Chromobacterium can easily be identified by hybridization with a 3H-labelled 23S rRNA probe from C. violaceum NCTC 9757: only members of C. violaceum have Tm(e) values between 76.5 and 80°C. [Chromobacterium] fluviatile strains have Tm(e) values of 70 to 71°C. Members of Janthinobacterium have Tm(e) values of 67 to 68°C and can further be identified by hybridization with a 3H-labelled 23S rRNA probe of J. lividum NCTC 9796. All other taxa are below 72°C Tm(e).

Special Features and Applications In this part, we call attention to products or metabolic pathways that have been intensively studied in C. violaceum and their possible applications. First of all, the biosynthesis of violacein (Riveros et al., 1988) and the different effects of this pigment have been studied. Violacein has antibiotic activity (DeMoss, 1967; Hoshino et al., 1987) and protects against predation by protozoa. In the older literature (Burbanck, 1942; Singh,

CHAPTER 3.2.8

The Genera Chromobacterium and Janthinobacterium

1942, 1945), C. violaceum and violacein itself are described as lethal to some ciliated protozoa. Amoebae and flagellates can also be killed by these bacteria (Singh, 1942). Later Groscop and Brent (1964) showed that violacein is toxic to soil amoebae while C. violaceum is toxic to Vorticella microstoma (Curds and Vandyke, 1966). Violacein has also a trypanocide effect (Riveros et al., 1988). The production of violacein can be used as a microbiological assay for L-tryptophan (Sebek, 1965). Apart from violacein, C. violaceum produces other antibiotics. Several were isolated and characterized in the last years: aerocyanidin, bearing an isonitrile group and being active against Gram-positive bacteria (Parker et al., 1988); aerocavin, exhibiting in vitro activity against both Gram-negative and Gram-positive bacteria (Singh et al., 1988); 3,6-dihydroxyindoxazene, inhibiting growth of Gram-negative bacteria (Hamada et al., 1983); and factor Y-T0678H (6hydroxy-3-oxo-1,2-benzisoxazolin), exhibiting a selective activity against Gram-negative bacteria (Imai et al., 1983). Other important compounds are produced by C. violaceum: arphamenines A and B, two specific inhibitors of aminopeptidase B that enhance immune responses (Umezawa et al., 1983). The production and function of C. violaceum aminopeptidases involved in enkephalin hydrolysis was studied by Weiss et al. (1983). Cooper et al. (1985) studied novel potentiators of β-lactam antibiotics and isolated two new compounds, SQ 28,504 and SQ 28,546, from a C. violaceum strain. These novel bacterial metabolites are glycopeptides which act synergistically with β-lactam antibiotics against Gram-negative bacteria. Earlier β-lactamase, primarily active against cephalosporins, was shown in a C. violaceum strain (Farrar and O’Dell, 1976). C. violaceum has also been used in studies on the production of HCN (Brysk et al., 1969; Niven et al., 1975), of unusual sugar compounds (Stevens et al., 1963), and of extracellular polysaccharides (Corpe, 1964). The tough membranous aspect of some C. violaceum colonies is the result of the latter polysaccharides (Corpe, 1964). Their role in the formation of soil aggregates has been suggested (Martin and Richards, 1963). The Copper-containing phenylalanine monooxygenase (Benkovic et al., 1986; Pember et al., 1987a, 1987b; Fujisawa and Nakata, 1987) and the denitrification processes (Bazylinski et al., 1986) were intensively studied. It was shown that denitrification terminates with the production of N2O and that NO2− reduction to N2O is not coupled to growth but may serve as a detoxification mechanism.

741

A C. violaceum strain has been isolated that can grow on and use acetonitrile as sole carbon source (K. D. Chapatwale et al., 1988) Another C. violaceum strain solubilized a considerable amount of pure gold after 40 days incubation in nutrient medium because the cyanide produced during growth and stationary phase formed the complex anion [Au(CN)2]− with gold (Smith and Hunt, 1985).

Janthinobacterium The basic description of this genus was given by Sneath in 1984. Janthinobacterium consists of Gram-negative rods with rounded ends, sometimes slightly curved. No resting stages are formed. They are usually motile by means of both a single polar flagellum and by one to four lateral flagella. They are strict aerobes. They produce violet colonies on solid media; in nutrient broth a violet ring is formed at the junction of the liquid surface and the container wall. They are resistant to benzylpenicillin (10 µg/ml) and to vibriostatic agent 0/129 (30 µg/disc). The most striking features are: 1) janthinobacteria are aerobic chemoorganotrophs with a strictly respiratory type of metabolism with oxygen as the terminal electron acceptor. 2) They have an optimum growth temperature of 25°C, a maximum growth temperature of 32°C and a minimum growth temperature of 2°C. 3) Only a few strains produce HCN. 4) They can grow rapidly on a medium containing citrate and ammonia as sole carbon and nitrogen source. 5) Nitrate and nitrite are reduced mostly with visible gas production. 6) The GC content is 61 to 67 mol%. They are soil and water organisms (Moss et al., 1978; Koburger and May, 1982) common in temperate climates. They are sometimes the causative agent of food spoilage. The type and sole species is J. lividum. Within this species, percentages of DNA binding of at least 48% were encountered between representative strains (De Ley et al., 1978). The membraneforming strains, formerly named C. amethystinum or C. membranaceum and sometimes referred to as “atypical C. lividum strains” (Moss and Ryall, 1981), and an unusual strain (ATCC 14487) which preferentially grows at 4°C all belong in J. lividum (De Ley et al., 1978; Sneath, 1984). The typical features in which Janthinobacterium differs from Chromobacterium can be found in Table 1.

Habitats J. lividum strains have been isolated from water and soil from rivers, creeks, lakes, and springs (Ryall and Moss, 1975; Koburger and May, 1982; Williams and Albright, 1984; Quevedo-

742

M. Gillis and J. de Ley

Sarmiento et al., 1986; Gonzalez et al., 1987). They mostly comprise only a small percentage of the total microflora, e.g., a mean of 5% of the total aerobic heterotrophic bacterial population in natural spring water (Quevedo-Sarmiento et al., 1986); a mean of 915 J. lividum isolates in 1 g water or soil (Koburger and May, 1982); and less than 1 to 3% of the total population sampled in lake water (Williams and Albright, 1984). According to Moss et al. (1978) they constitute approximately 75% of the total amount of purple-pigmented bacteria isolated from water and soil samples in a lowland river (river Wey in the UK). They are frequently isolated from a variety of foods, i.e., processed poultry (Cox, 1975); turbot (Mudarris and Austin, 1988); various vegetables (Koburger and May, 1982); and Japanese noodles (Naito et al., 1986a). In turbot gills (Mudarris and Austin, 1988), J. lividum represented approximately one-third of the aerobic heterotrophic bacteria (7 × 105/g wet weight of gill tissue). In vegetables, they are present in quantities varying from 130 to over 6000/g (Koburger and May, 1982). Isolates from slaughterhouse effluents (Etherington et al., 1976) were colorless, could grow on meat, and exhibited a high proteolytic activity (Dainty et al., 1978), which is exceptional for members of this genus (Sneath, 1984). Bettelheim et al. (1968) isolated bacterial strains from germinating seeds of Psychotria nairobiensis (Rubiaceae) and Ardisia crispa (Myrsinaceae), which they identified then as C. lividum, thus now J. lividum. By an immunofluorescence technique, Bettelheim et al. (1968) detected these organisms in tissues of several leaf-nodulated Rubiaceae and Myrsinaceae, and presented the hypothesis that some J. lividum strains are bacterial endophytes. We shall not review the discussions and disagreements in the literature on this issue (Lersten and Horner, 1976; see also The Genera Chromobacterium and Janthinobacterium in Volume 5) because it was shown that Bettelheim’s isolates are not genuine janthinobacteria (De Ley et al., 1978) since they belong in rRNA superfamily IV, where they are a member of the [Flavobacterium] capsulatum rRNA branch (The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition). The role, if any, of J. lividum strains in the induction of sporangia in Phytophthora cinnamomi has been discussed (Zentmeyer, 1965; Broadbent et al., 1974).

Enrichment and Isolation As mentioned before, Janthinobacterium can be enriched and isolated on the same media as given in detail for Chromobacterium above.

CHAPTER 3.2.8

According to Moss and Ryall (1981), they grow rapidly and can be selected on a citrate ammonium salts agar medium: Citric acid NaCl MgSO4 ·7H 2O NH4H2PO4 K2HPO4 Ionoagar nr. 2 (Oxoid) Distilled water

2g 1g 0.2 g 1g 1g 15 g 1 liter

Koburger and May (1982) recovered J. lividum from a variety of food samples only on Bennett’s agar (see above) at 25°C. On this medium they found higher counts in water and soil samples than on the medium of Ryall and Moss (1975) or on Aeromonas membrane agar (Rippey and Cabelli, 1979). Maximal numbers of J. lividum were isolated at 25°C from turbot gills (Mudarris and Austin, 1988) on the following medium (%w/v or v/v): CaCl2 Yeast extract Beef extraxt Casein Tryptone Agar no. 1 Aged sea water pH

0.1 0.1 0.5 0.6 0.2 1.5 75 7.2

Identification Characteristics differentiating J. lividum from C. violaceum and Iodobacter fluviatilis* have been given in Table 1. Differences with Alteromonas luteoviolacea have been reported above. Although colorless J. lividum strains could be confused with Pseudomonas fragi (Moss and Ryall, 1981), Dainty et al. (1978) were able to identify correctly their nonpigmented isolates as J. lividum. Because J. lividum strains constitute a narrow rRNA cluster with Tm(e) values between 77 and 79°C vs. 3H labelled 23S rRNA from J. lividum NCTC 9796, they can easily be identified by hybridizing with this rRNA probe. [Pseudomonas] rubrisubalbicans, [Aquaspirillum] autotrophicum, and an as yet unnamed group of strains isolated from clinical material and described by E. Falsen (Culture Collection, University of Göteborg, Sweden) have Tm(e) values of approximately 76°C and are the closest neighbors of J. lividum (see The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition). Phenotypically (M. Goor et al. 1986; see The Genus Aquaspirillum in Volume 5) it is possible to differentiate J. *The latter species was originally named Iodobacter fluviatile, but since Iodobacter is masculine, it obviously has to be Iodobacter fluviatilis.

CHAPTER 3.2.8

The Genera Chromobacterium and Janthinobacterium

743

Table 2. Characteristics differentiating Janthinobacterium from [Pseudomonas] rubrisubalbicans and similar unnamed clinical isolates. Character Urease Hydrolysis of esculin Growth on D-fucose, 5-keto-gluconate, acetate, caprate, amylamine, and tryptamine Growth on L-histidine

Janthinobacterium

[Pseudomonas] rubrisubalbicans and unnamed clinical isolatesa

− + −

+ − +

+

−

Isolates of Falsen (1989).

lividum from the group formed by [Pseudomonas] rubrisubalbicans and the unnamed clinical isolates (Table 2) and from [Aquaspirillum] autotrophicum (see Table 1 in The Genus Aquaspirillum in Volume 5). Results from M. Goor et al. (1986). Preservation of Strains J. lividum strains survive for several years at 4°C in 0.1% peptone or at −80°C in nutrient broth with 15% glycerol as a cryoprotectant. Lyophilization and storage at 4°C is also applied although the viability may be low (Sneath, 1984).

Special Features and Applications For the different effects of violacein, we refer to the discussion under “Chromobacterium” above. The antimicrobial activity of violacein isolated from J. lividum has recently been described (Naito et al., 1986b). The location of the pigment was investigated by Lin and Kester (1988), and in broth cultures, both intracellular and extracellular pigment was found. Intracellular pigment is located in the cell membrane. Extracellular pigment is water soluble and shows a blue shift when compared to solvent-extracted pigment. It was suggested that extracellular pigment is possibly non-covalently bound to a small protein. J. lividum produces 2,5-diketogluconate (Bernaerts and De Ley, 1971) via 2-ketogluconate. Kwok et al. (1987) described a J. lividum strain as a bacterial antagonist inducing suppression of Rhizoctonia damping-off in container media amended with composted hardwood tree bark. Strains from food produce sometimes extracellular metallo-proteinases (Etherington et al., 1976; Dainty et al., 1978) which could play a role in tenderizing meat. A J. lividum strain was used to study the effects of the insecticide acephate on bacterial growth and nutrient uptake (Williams and Albright, 1984). Acknowledgments. We are indebted to the National Fund for Scientific Research (Belgium) and the Fund for Medical Scientific Research (Belgium) for research and personnel grants.

Literature Cited Aldridge, K. E., G. T. Valainis, C. V. Sanders. 1988. Comparison of the in-vitro activity of ciprofloxacin and 24 other antimicrobial agents against clinical strains of Chromobacterium violaceum. Diagn. Microbiol. Infect. Dis. 10:31– 40. Bazylinski, D. A., E. Palome, N. A. Blakemore, R. P. Blakemore. 1986. Denitrification by Chromobacterium violaceum. Appl. Environm. Microbiol. 52:696–699. Benkovic, S. J., L. M. Bloom, G. Bollag, T. A. Dix, B. J. Gaffney, S. Pember. 1986. The mechanism of action of phenylalanine hydroxylase. Ann. N.Y. Acad. Sci. 471:226–232. Bernaerts, M., J. De Ley. 1971. 2,5-Diketogluconate formation by Chromobacterium. J. Microbiol. Serol. 37:185– 195. Bettelheim, K. A., J. F. Gordon, J. Taylor. 1968. The detection of a strain of Chromobacterium lividum in the tissues of certain leaf nodulated plants by the immunofluorescence technique. J. Gen. Microbiol. 54:177–184. Broadbent, P., K. F. Baker, J. Aust. 1974. Association of bacteria with sporangium formation and breakdown of sporangia in Phytophthora sp. Austr. J. Agricultural Research 25:139–145. Brysk, M. M., W. A. Corpe, L. V. Hankes. 1969. β-Cyanoalanine formation by Chromobacterium violaceum. J. Bacteriol. 97:322–327. Buchanan, R. E. 1918. Studies in the nomenclature and classification of bacteria. J. Bacteriol. 3:27–61. Burbanck, W. D. 1942. Physiology of the ciliate Colpidium colpoda. Physiological Zoology 15:342–362. Chapatwala, K. D., J. D. Richardson, M. Nawaz, J. H. Wolfram. 1988. Isolation and identification of acetonitrile degrading bacteria. Abstr. Annual Meeting American Society Microbiology, May 8–13, Miami Beach, Florida, USA. 88:225. Collins, B., G. Kolias, A. Battles, A. Moreland. 1985. Chromobacterium violaceum infection in a stumptailed monkey Macaca arctoides. Lab. Anim. Sci. 35:530–531. Cooper, R., J. S. Wells, R. B. Sykes. 1985. Novel potentiators of β-lactam antibiotics. J. Antibiot. 38:449–454. Corpe, W. A. 1951. A study of the wide spread distribution of Chromobacterium species in soil by a simple technique. J. Bacteriol. 62:515–517. Corpe, W. A. 1964. Factors influencing growth and polysaccharide formation by strains of Chromobacterium violaceum. J. Bacteriol. 88:1433–1441. Cox, N. A. 1975. Isolation and identification of a genus, Chromobacterium, not previously found on processed poultry. Appl. Microbiol. 29:864.

744

M. Gillis and J. de Ley

Curds, C. R., J. M. Vandyke. 1966. The feeding habits and growth rates of some fresh-water ciliates found in activated sludge plants. J. Appl. Ecology 3:127–137. Dainty, R. H., D. J. Etherington, B. G. Shaw, J. Barlow, G. T. Banks. 1978. Studies on the production of extracellular proteinases by a non-pigmented strain of Chromobacterium lividum isolated from abattoir effluent. J. Appl. Bacteriol. 45:111–124. De Ley, J., P. Segers, M. Gillis. 1978. Intra-and intergeneric similarities of Chromobacterium and Janthinobacterium ribosomal ribonucleic acid cistrons. Int. J. Syst. Bacteriol. 28:154–168. DeMoss, R. D. 1967. Violacein. 77–81.D. Gottlieb, and P. Shaw (ed.) Mechanisms of action and biosynthesis of antibiotics, vol. 2. Springer-Verlag. New York. Dewhirst, F. E., B. J. Paster, P. L. Bright. 1989. Chromobacterium, Eikenella, Kingella, Neisseria, Simonsiella and Vitreoscilla species comprise a major branch of the beta group Proteobacteria by 16S ribosomal ribonucleic acid sequence comparison: Transfer of Eikenella and Simonsiella to the family Neisseriaceae (emend.). Int. J. Syst. Bacteriol. 39:258–266. Etherington, D. J., P. B. Newman, R. H. Dainty, S. M. Partridge. 1976. Purification and properties of the extracellular metallo-proteinases of Chromobacterium lividum (NCIB 10926). Biochim. Biophys. Acta 445:739–752. Falsen, E. 1989. Catalogue of strains, Culture Collection. University of Göteborg. Göteborg, Sweden. Farrar, W. E., N. M. O’Dell. 1976. β-Lactamase activity in Chromobacterium violaceum. J. Infect. Dis. 134:290– 293. Feldman, R. B., G. A. Stem, C. I. Hood. 1984. Chromobacterium violaceum infection of the eye. Arch. Ophthalmol. 102:711–713. Fujisawa, H., H. Nakata. 1987. Phenylalanine 4-monooxygenase from Chromobacterium violaceum. Methods Enzymol. 142:44–49. Gauthier, M. J. 1976. Morphological, physiological and biochemical characteristics of some violet-pigmented bacteria isolated from seawater. Can. J. Microbiol. 22: 138–149. Gauthier, M. J. 1982. Validation of the name Alteromonas luteoviolacea. Int. J. Syst. Bacteriol. 32:82–86. Gogolewski, R. P. 1983. Chromobacterium violaceum septicaemia in a dog. Aust. Vet. J. 60:226. Gonzalez, C., C. Gutierrez, T. Grande. 1987. Bacterial flora in bottled uncarbonated mineral drinking water. Can. J. Microbiol. 33:1120–1125. Goor, M., E. Falsen, B. Pot, M. Gillis, K. Kersters, J. De Ley. 1986. Taxonomic position of the phytopathogen Pseudomonas rubrisubalbicans and related clinical isolates. 37.Abstr. XIV International Congress of Microbiology, 7–13 September, Manchester, UK. Groscop, J. A., M. N. Brent. 1964. The effects of selected strains of pigmented microorganisms on small freeliving amoebae. Can. J. Microbiol. 10:579–584. Groves, M. G., J. M. Strauss, J. Abbas, C. E. Davis. 1969. Natural infections of gibbons with a bacterium producing a violet pigment (Chromobacterium violaceum). J. Inf. Diseases 120:605–610. Hamada, M., S. Kondo, H. Nakamura, T. Ikeda, D. Ikeda, K. Iinuma, S. Gomi, Y. Ikeda, T. Takeuchi, H. Umezawa, Y. Iitaka. 1983. A new antibiotic, 3,6-dihydroxyindoxazene. J. Antibiot. 36:445–447. Hoshino, T., T. Kondo, T. Uchiyama, N. Ogasawara. 1987. Biosynthesis of violacein a novel rearrangement in tryp-

CHAPTER 3.2.8 tophan metabolism with a 1 2-shift of the indole ring. Agric. Biol. Chem. 51:965–968. Hussain, A., V. Van cˇ ura. 1970. Formation of biologically active substances by rhizosphere bacteria and their effect on plant growth. Folia Microbiologica 15:468– 478. Imai, H., K. Suzuki, S. Miyazaki, K. Tanaka, S. Watanabe, M. Iwanami. 1983. A new antibiotic Y-TO678H produced by a Chromobacterium species. J. Antibiot. 36:911–912. Johnson, E. A., R. J. S. Beer. 1971. Violacein, spectrum no. J8/2. H. M. Perkampus, I. Sandeman, and C. J. Timmons (ed.) UV atlas of organic compounds, vol. 5. Butterworth. London. Johnson, W. M., A. F. Disalvo, R. R. Steuer. 1971. Fatal Chromobacterium violaceum septicemia. Am. J. Clin. Pathol. 56:400– 406. Kaufman, S. C., D. Ceraso, A. Schugurensky. 1986. First case report from Argentina of fatal septicemia caused by Chromobacterium violaceum. J. Clin. Microbiol. 23:956– 958. Keeble, J. R., T. Cross. 1977. An improved medium for the enumeration of Chromobacterium in soil and water. J. Appl. Bacteriol. 43:325–327. Koburger, J. A., S. O. May. 1982. Isolation of Chromobacterium spp. from foods, soil, and water. Appl. Environm. Microbiol. 44:1463–1465. Kwok, O. C. H., P. C. Fahy, H. A. J. Hoitink, G. A. Kuter. 1987. Interactions between bacteria and Trichoderma hamatum in suppression of rhizoctonia damping-off in bark compost media. Phytopathology 77:1206–1212. Laws, L., W. T. R. Hall. 1964. Chromobacterium violaceum infections in a pig. Queensland J. Agric. Sci. 20:393– 400. Lersten, N. R., H. T. Horner, Jr. 1976. Bacterial leaf nodule symbiosis in angiosperms with emphasis on Rubiaceae and Myrsinaceae. Bot. Rev. 42:145–214. Lin, Y. C., A. S. Kester. 1988. Production, location and binding of violacein in Janthinobacterium. Abstr. Annual Meeting of the American Society for Microbiology, Miami Beach, FL. 88:186. Liu, C. H., C. N. Weng, Y. L. Lin, R. M. Chu. 1988. Pathology of experimental infection with Chromobacterium violaceum in pigs. J. Clin. Soc. Vet. Sci. 14:191–202. Logan, N. A. 1989. Numerical taxonomy of violet-pigmented, Gram-negative bacteria and description of Iodobacter fluviatile gen. nov., com. nov. Int. J. Syst. Bacteriol. 39:450– 456. Machin Villafranca, C., M. Ley, L. Torres Hernandez. 1986. Infeccion por Chromobacterium violaceum. Rev. Cubana Med. Trop. 38:353–357. Martin, J. P., S. J. Richards. 1963. Decomposition and binding action of polysaccharide from Chromobacterium violaceum in soil. J. Bacteriol. 85:1288–1294. McClure, H. M., J. Chang. 1976. Chromobacterium violaceum infection in a nonhuman primate (Macaca assamensis). Lab. Anim. Sci. 26:807–810. Miller, D. P., W. T. Blevins, D. B. Steele, M. D. Stowers. 1988. A comparative study of virulent and avirulent strains of Chromobacterium violaceum. Can. J. Microbiol. 34:249– 255. Moffet, M. L., R. R. Colwell. 1968. Adansonian analysis of the Rhizobiaceae. J. Gen. Microbiol. 51:245–266. Moss, M. O., T. N. Bryant. 1982. DNA:rRNA hybridization studies of Chromobacterium fluviatile. J. Gen. Microbiol. 128:829–834. Moss, M. O., C. Ryall. 1981. The genus Chromobacterium. 1355–1364.M. P. Starr, H. Stolps, H. G. Trü per, A. B. Balows, and H. G. Schlegel (ed.) The prokaryotes: A

CHAPTER 3.2.8

The Genera Chromobacterium and Janthinobacterium

handbook in habitats, isolation, and identification of bacteria. Springer-Veriag. Berlin. Moss, M. O., C. Ryall, N. A. Logan. 1978. The classification and characterization of chromobacteria from a lowland river. J. Gen. Microbiol. 105:11–21. Mudarris, M., B. Austin. 1988. Quantitative and qualitative studies of the bacterial microflora of turbot Scophthalmus maximus L. gills. J. Fish. Biol. 32:223–229. Naito, S., I. Shiga, N. Yamaguchi. 1986a. Isolation and identification of violet pigment producing bacteria from yudemen boiled japanese noodle and protection against microbiological deterioration. Part X. J. Jap. Soc. Food Sci. Technol. 33:752–758. Naito, S., I. Shiga, N. Yamaguchi. 1986b. Antimicrobial activity of violet pigment produced by Janthinobacterium lividum isolated from yudemen boiled japanese noodle. Part XIV. J. Jap. Soc. Food Sci. Technol. 33:759–763. Niven, D. F., P. A. Collins, C. J. Knowles. 1975. The respiratory system of Chromobacterium violaceum grown under conditions of high and low cyanide evolution. J. Gen. Microbiol. 90:271–285. Ognibene, A. J., E. Thomas. 1970. Fatal infection due to Chromobacterium violaceum in Vietnam. Am. J. Clin. Pathol. 54:607–610. Onile, A., B. O. Sobowale, T. Odugbemi. 1984. Human infection due to Chromobacterium violaceum: a report from Ilorin, Nigeria. East Afr. Med. J. 61:849–852. Parker, W. L., M. L. Rathnum, J. H. Johnson, J. S. Wells, P. A. Principe, R. B. Sykes. 1988. Aerocyanidin, a new antibiotic produced by Chromobacterium violaceum. J. Antibiot. 41:454– 460. Pember, S. O., J. J. Villafranca, S. J. Benkovic. 1987a. Chromobacterium violaceum phenylalanine 4-monooxygenase. Methods Enzymol. 142:50–56. Pember, S. O., S. J. Benkovic, J. J. Villafranca, M. Passenkiewicz-Gierula, W. E. Antholine. 1987b. Adduct formation between the cupric site of phenylalanine hydroxylase from Chromobacterium violaceum and 6,7-dimethyltetrahydropterin. Biochemistry 26:4477– 4483. Petrillo, V. F., V. Severo, M. M. Santos, E. L. Edelweiss. 1984. Recurrent infection with Chromobacterium violaceum: first case report from South America. J. Infect. 9:167– 169. Quevedo-Sarmiento, J., A. Ramos-Cormenzana, J. GonzalezLopez. 1986. Isolation and characterization of aerobic heterotrophic bacteria from natural spring waters in the Lanjaron area (Spain). J. Appl. Bacteriol. 61:365–372. Rippey, S. R., V. J. Cabelli. 1979. Membrane filter procedure for enumeration of Aeromonas hydrophila in fresh waters. Appl. Environm. Microbiol. 38:108–113. Riveros, R., M. Haun, V. Campos, N. Duran. 1988. Bacterial chemistry IV. Arq. Biol. Techol. (Curitiba) 31:475– 487. Rossau, R., G. Vandenbussche, S. Thielemans, P. Segers, H. Grosch, E. Göthe, W. Mannheim, J. De Ley. 1989. Ribosomal ribonucleic nucleic acid cistron similarities and deoxyribonucleic acid homologies of Neisseria, Kingella, Eikenella, Simonsiella, Alysiella, and Centers for Disease Control groups EF-4 and M-5 in the emended family Neisseriaceae. Int. J. Syst. Bacteriol. 39:185–198. Ryall, C., M. O. Moss. 1975. Selective media for the enumeration of Chromobacterium ssp. in soil and water. J. Appl. Bacteriol. 38:53–59. Sebek, O. K. 1965. Microbiological method for the determination of L-tryptophan. J. Bacteriol. 90:1026–1031.

745

Shetty, M., A. Venkatesh, S. Shenoy, P. G. Shivananda. 1987. Chromobacterium violaceum meningitis: a case report. Indian J. Med. Sci. 41:275–276. Simo, F., P. D. Reuman, F. J. Martinez, E. M. Ayoub. 1984. Chromobacterium violaceum as a cause of periorbital cellulitis. Pediatr. Infect. Dis. 3:561–563. Singh, B. N. 1942. Toxic effects of certain bacterial metabolic products on soil protozoa. Nature (London) 149:168. Singh, B. N. 1945. The selection of bacterial food by soil amoebae, and the toxic effects of bacterial pigments and other products on soil protozoa. Br. J. Exp. Pathol. 26:316–325. Singh, P. D., W.-C. Liu, J. Z. Gougoutas, M. F. Malley, M. A. Porubcan, W. H. Trejo, J. S. Wells, R. B. Sykes. 1988. Aerocavin, a new antibiotic produced by Chromobacterium violaceum. J. Antibiot. 41:446–453. Sivendra, R., S. H. Tan. 1977. Pathogenicity of nonpigmented cultures of Chromobacterium violaceum. J. Clin. Microbiol. 5:514–516. Smith, A. D., R. J. Hunt. 1985. Solubilization of gold by Chromobacterium violaceum. J. Chem. Technol. Biotechnol. 35:110–116. Sneath, P. H. A. 1956. Cultural and biochemical characteristics of the genus Chromobacterium. J. Gen. Microbiol. 15:70–98. Sneath, P. H. A. 1960. A study of the bacterial genus Chromobacterium. Iowa State J. Sci. 34:243–500. Sneath, P. H. A. 1974. Chromobacterium Bergonzini 1881. 354–357.R. E. Buchanan, and N. E. Gibbons (ed.) Bergey’s manual of determinative bacteriology. Williams and Wilkins. Baltimore. Sneath, P. H. A. 1984. Chromobacterium Bergonzini 1881. 580–582.N. R. Krieg, and J. G. Holt (ed.) Bergey’s manual of systematic bacteriology, vol. 1. Williams and Wilkins. Baltimore. Sneath, P. H. A., J. P. F. Whelan, R. B. Singh, D. Edwards. 1953. Fatal infection by Chromobacterium violaceum. Lancet ii:276–277. Sorensen, R. U., M. R. Jacobs, S. B. Shurin. 1985. Chromobacterium violaceum adenitis acquired in the northern United States as a complication of chronic granulomatous disease. Pediatr. Infect. Dis. 4:701–702. Stackebrandt, E., R. G. E. Murray, H. G. Trü per. 1988. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “Purple bacteria and their relatives.” Int. J. Syst. Bacteriol. 38:321–325. Stevens, C. L., P. Blumbergs, F. A. Daniher, R. W. Wheat, A. Kujomoto, E. L. Rollins. 1963. The identification and synthesis of the 4-amino sugar from Chromobacterium violaceum. J. Am. Chem. Soc. 85:3061. Suarez, A. E., B. Wenokur, J. M. Johnson, L. D. Saravolatz. 1986. Nonfatal chromobacterial sepsis. South. Med. J. 79:1146–1148. Umezawa, H., T. Aoyagi, S. Ohuchi, A. Okuyama, H. Suda, T. Takita, M. Hamada, T. Takeuchi. 1983. Arphamenines A and B, new inhibitors of aminopeptidase B, produced by bacteria. J. Antibiot. 36:1572–1575. Van Landschoot, A., J. De Ley. 1983. Intra- and intergeneric similarities of the rRNA cistrons of Alteromonas, Marinomonas (gen. nov.) and some other Gram-negative bacteria. J. Gen. Microbiol. 129:3057–3074. Victoria, B., H. Baer, E. M. Ayoub. 1974. Successful treatment of systemic Chromobacterium violaceum infection. J. Am. Med. Assoc. 230:578–580. Weiss, B., K. S. Hui, M. Hui, A. Lajtha. 1983. Effect of bestatin analogues and other compounds on enkephalin

746

M. Gillis and J. de Ley

hydrolysis by an aminopeptidase from the mesophiles Pseudomonas sp. ATCC 11299A and Chromobacterium violaceum ATCC 12540. Res. Commun. Chem. Pathol. Pharmacol. 39:463– 475. Wilkey, I. S., A. McDonald. 1983. A probable case of Chromobacterium violaceum infection in Australia. Med. J. Aust. 2:39–40. Williams, G. L., L. J. Albright. 1984. The effects of the insecticide acephate on the growth and nutrient uptake of an aquatic bacterium. Can. J. Microbiol. 30:375–380. Woese, C. R., W. G. Weisburg, B. J. Paster, C. M. Hahn, R. S. Tanner, N. R. Krieg, H.-P. Koops, H. Harms, E. Stacke-

CHAPTER 3.2.8 brandt. 1984. The phylogeny of purple bacteria: the beta subdivision. Syst. Appl. Microbiol. 5:327–336. Wu, S. H., S. J. Lin, H. M. Tso, C. B. Liu, W. C. Tsai. 1986. (Fatal septicemia due to Chromobacterium violaceum). Chung Hua Min Kuo Wei Sheng Wu Chi Mien I Hsueh Tsa Chic 19:289–294. Zentmyer, G. A. 1965. Bacterial stimulation of sporangium production in Phytophthora cinnamomi. Science 150:1178–1179. Zwang, J., Z. G. Mai, Z. H. Dung. 1987. Mobilization of granulocyte-macrophage colony forming cell CFU. Acta Physiol. Sin. 39:61–67.

Prokaryotes (2006) 5:747–750 DOI: 10.1007/0-387-30745-1_33

CHAPTER 3.2.9 ehT

areneG

mu i re t cabo l l yhP

dna

mur t caborhcO

The Genera Phyllobacterium and Ochrobactrum JEAN SWINGS, BART LAMBERT, KAREL KERSTERS AND BARRY HOLMES

The study of leaf nodulation in several tropical plant families dates back to the beginning of the 20th century. Various genera of bacteria, e.g., Phyllobacterium, were thought to be associated with leaf nodules (see below). Phyllobacterium was described by Knösel (1962) but the genus only found wide taxonomic recognition with the publication of Bergey’s Manual of Systematic Bacteriology (Knösel, 1984). DNArRNA hybridizations revealed a close relationship between Phyllobacterium and bacteria from the Centers for Disease Control (CDC) group Vd within the “alpha” subclass (rRNA superfamily IV) of the proteobacteria together with those throughout of the genera Brucella, Rhizobium, Mycoplana, and Agrobacterium (De Ley et al., 1987; Stackebrandt et al., 1988). Holmes et al. 1988 proposed the name Ochrobactrum anthropi for CDC group Vd. They also demonstrated that Phyllobacterium showed closest phenotypic similarity to Ochrobactrum anthropi. In the present chapter, the genera Phyllobacterium and Ochrobactrum are treated together because of their close phylogenetic relatedness. Nevertheless, the ecological niches from which these bacteria have been isolated are quite different: Phyllobacterium strains occur in leaf nodules and in the rhizosphere, whereas Ochrobactrum strains have been isolated almost exclusively from human clinical material. In clinical microbiology, Ochrobactrum, more recently described as Achromobacter group Vd until formally named, was usually treated together with Achromobacter xylosoxidans (more recently known as Alcaligenes xylosoxidans subsp. xylosoxidans) (Gilardi, 1978; Rubin et al., 1985). It has now become apparent that Ochrobactrum and Phyllobacterium are not related phylogenetically to Achromobacter xylosoxidans, which belongs to the Alcaligenaceae in the “beta” subclass (rRNA superfamily III) of the proteobacteria (De Ley et al., 1986).

Habitats The Occurrence of Phyllobacterium in Leaf Nodules Plants of over 400 species of the families Rubiaceae and Myrsinaceae are reported to produce leaf nodules. Short-lived bacterial colonies become established intercellularly and die before full leaf expansion (Lersten and Horner,

This chapter was taken unchanged from the second edition.

1976). Bacterial leaf nodules were defined by Horner and Lersten (1972) as: Internal cavities in the leaf lamina, open to the exterior by way of stomatal pores only in early stages of nodule development. The bacteria inhabiting such nodules are part of a population maintained by the host plant in the vegetative and floral buds and passed to the succeeding generations through the seeds. This definition rules out all casual bacteriaplant associations. Despite more than a century of research on leaf nodulation (for a review, see Lersten and Horner, 1976) it is not yet clear which bacteria are involved and how they participate in the nodulation process. In the past, the bacterial symbionts in the nodules were recognized as belonging to the genera Chromobacterium, Xanthomonas, Klebsiella and Phyllobacterium (Horner and Lersten, 1972). Knösel (1962) isolated and described Phyllobacterium and stated that it is able to induce characteristic nodules on leaves (Knösel, 1984), basing his conclusions on earlier investigations by Miehe (1911), von Faber (1912), and de Jongh (1938). However, it is not clear whether the bacteria previously studied by these workers are identical to those described by Knösel. There is no proof for the leaf nodulation capacity of Phyllobacterium.

The Occurrence of Phyllobacterium in the Rhizosphere Pseudomonas fluorescens and Phyllobacterium were found to be the two most frequently occurring bacteria on root surfaces during a large-scale assessment of the rhizobacterial communities of young sugar beet plants (between the second and 10th leaf stage) (Lambert et al., 1990). These bacteria were found in 198 out of 1,100 plants investigated in Belgium and Spain at densities up to 2 × 108 per gram of root. This was the first record of the occurrence of Phyllobacterium in the rhizosphere. Extensive analyses of the microflora from the rhizoplane of other crop plants have not revealed the presence of Phyllobacterium.

748

J. Swings et al.

Occurrence of Ochrobactrum (CDC group Vd) in Clinical Samples and the Environment Ochrobactrum anthropi strains have been isolated predominantly from human blood, urogenital tracts, urine, respiratory tracts, ears, and wounds (Tatum et al., 1974; Holmes et al., 1988). Only a few strains were isolated from feces, spinal fluid, eye, hospital apparatus, or the environment (arsenical cattle-dipping fluid, soil, sewage, and from thin-layer Sephadex plates) (Holmes et al., 1988). The clinical significance of O. anthropi remains largely unknown; strains of this species are considered to be opportunistic pathogens. One strain has been reported in association with a pancreatic abscess (Appelbaum and Campbell, 1980) and another has been associated with a puncture wound leading to osteochondritis of a foot (Barson et al., 1987). Isolation Isolation of Phyllobacterium from the Sugar Beet Rhizosphere (Lambert et al., 1990) The entire root system of a plant is carefully washed to remove adhering soil, then vigorously shaken for 15 min using a flask shaker in a phosphate buffered saline solution containing 0.025% Tween 20. Serial dilutions of the resulting suspensions are plated on 10% trypticase soy broth plus 2% agar (TSBA). After incubation at 28°C for 2 days, Phyllobacterium colonies are beige-colored, large (±5 mm), and convex with an entire edge. They are very mucoid and glistening. Several subcultures are necessary in order to obtain pure isolates. Phyllobacterium readily grows on other nonselective media for aerobic, heterotrophic bacteria (nutrient agar, trypticase soy agar, Luria-Bertani agar, etc.).

Isolation of Phyllobacterium from Leaf Nodules (Knösel, 1984) Washed leaf pieces containing nodules are macerated by rubbing and placed in saline. After shaking, serial dilutions of the suspension are plated onto carrot juice agar containing yeast extract. After incubation at 28°C, a variety of pigmented and nonpigmented colonies develops. Typical colonies are transferred into liquid carrot juice medium which is then used to inoculate the tests used for identification. The cultures should be observed after 24 to 48 h to confirm the presence of star clusters.

Isolation of Ochrobactrum anthropi from Clinical Material Samples from human blood, urine and urogenital tract, sputum, wound, throat, stool, rectum, and other material are plated on blood agar or on MacConkey agar (Rubin et al., 1985; Holmes et al., 1988). Two colony types were recognized on blood agar at 35°C after 48 h: 1) an extremely mucoid type, 0.1 to 4 mm in diameter, opaque and gray-white, showing beta-hemolysis; and 2) a pinpoint type, opaque and semiglossy, gray or white, showing beta-hemolysis after prolonged incubation (Chester and Cooper, 1979). On MacConkey agar after continued incubation, colonies from 3 to 5 mm diameter were formed, opaque and pink to purple. Half the strains produced

CHAPTER 3.2.9 colonies with such notably gummy consistency that it was difficult to remove them from the agar surface (Chester and Cooper, 1979). On MacConkey agar, Ochrobactrum can be easily overlooked due to the small size of the colonies after overnight incubation (Chester and Cooper, 1979).

Preservation of Cultures Well-grown cultures may be kept at 4°C for 2 to 3 months on nutrient agar slants in screw-capped bottles. Isolates can be stored freeze-dried or as a suspension in 25% glycerol at −70°C.

Identification Phyllobacterium and Ochrobactrum are both Gram-negative, strictly aerobic bacteria with a respiratory type of metabolism. They show oxidase and catalase activity and both grow on nutrient agar. Indole is not produced. They oxidize glucose and xylose but not lactose (Clark et al., 1984; G. L. Gilardi, personal communication, Lambert et al., 1990). They do not produce extracellular enzymes for the hydrolysis of Tween 80, DNA, gelatin, starch, or casein (Gilardi, 1978). The API 20NE system will identify them as Achromobacter sp., which is still a “dumping ground” (Kersters and De Ley, 1984) for various aerobic peritrichously flagellated rods. Recent work on Phyllobacterium and Ochrobactrum has led to the differentiation scheme given in Table 1. An extensive description of Phyllobacterium and Ochrobactrum can be found in Lambert et al. (1990) and Holmes et al. (1988), respectively. A comment should be made on the flagellar arrangement of strains of both genera. Phyllobacterium and Ochrobactrum strains both show 1 to 3 polar, subpolar, or Table 1. Features that differentiate Phyllobacterium and Ochrobactrum (CDC group Vd). Property Utilizationa of: L-Citrulline Glutarate Erythritol Glycine L-Tryptophan Hydrolysisb of paranitrophenol-αmaltoside Hydrolysisb of paranitrophenol-αxylopyranoside

Phyllobacteriumc

Ochrobactrumd

− − − − + +

+ + + + − −

+

−

Using API 50CH and 50AO, 50AA strips (API System, La Balme-les-Grottes, Montalieu-Vercieu, France). Using API ZYM strips. 15 strains total. From Lambert et al. (1990). 56 strains total. From Holmes et al. (1988).

CHAPTER 3.2.9

The Genera Phyllobacterium and Ochrobactrum

lateral flagella (Lambert et al., 1990). According to Holmes et al. 1988 and Gilardi (1978) Ochrobactrum strains are characterized by peritrichous flagella, but other workers have found only polar, subpolar, or lateral flagella but no definitely peritrichous flagella (Clark et al., 1984; Chester and Cooper, 1979). An unambigous identification of Phyllobacterium or Ochrobactrum is possible by application of carbon assimilation tests (API 50CH, 50AO, or 50AA) or various conventional tests (Tatum et al., 1974; Rubin et al., 1985; Holmes et al., 1988; Lambert et al., 1990) or by protein electrophoretic fingerprinting. Variability in Gram reaction, the presence of curved cells with swollen-ends, and the formation of extremely small colonies after overnight incubation, which develop into mucoid colonies upon continued incubation, may lead to incorrect identification as Corynebacterium or Klebsiella (Chester and Cooper, 1979). Knösel (1984) used the criterium of nitrate reduction to differentiate Phyllobacterium myrsinacearum (+) from P. rubiacearum (−). Lambert et al. (1990), however, found that P. rubiacearum LMG 1t1T reduced nitrates. Using the API 2ONE system, they found that all Phyllobacterium strains showed a slight NO3− reduction. These observations cast serious doubt on the usefulness of this criterion for species differentiation within the genus Phyllobacterium. All Ochrobactrum strains are capable of nitrate reduction (Gilardi, 1978; Clark et al., 1984; Holmes et al., 1988).

al., 1990), which occur in all the classical nitrogen-fixing bacteria. Phyllobacterium is able to interact with plant tissues, as demonstrated by the tumor induction by Ti-plasmid-carrying Phyllobacterium strains on Kalanchoe plants (Lambert et al., 1990). Tumor induction involves the attachment of bacterial cell wall sites to plant cell wall sites prior to the induction of T-DNA transfer. The chromosomal genes chvA, chvB, exoC, and att are the only ones found to be involved in attachment but could not be detected in Phyllobacterium strains, suggesting that other genes are present with similar functions. There are no data indicating that Phyllobacterium is pathogenic or deleterious to plants. A striking feature, common to both Phyllobacterium and Ochrobactrum, is nutritional versatility, which makes their rapid growth and proliferation possible in rich environments such as the root surface and clinical sites. A number of Phyllobacterium isolates showed antifungal and antibacterial activities (Lambert et al., 1990). The oxidation of lactose to 3-ketolactose is a unique feature of many Agrobacterium strains (Kersters and De Ley, 1984) and is negative for all Phyllobacterium and Ochrobactrum strains.

Physiological Properties Cellular fatty acids of Ochrobactrum anthropi (Dees and Moss, 1978) as well as antimicrobial susceptibilities have been determined (von Graevenitz, 1985; Gilardi, 1989). Growth of Phyllobacterium strains was inhibited by doxycycline, novobiocin, framycetin, and tetracycline (Lambert et al., 1990). The most intriguing question that remains unanswered is whether Phyllobacterium has a real symbiotic cyclical relationship with its hosts. Reinfection studies did not allow unambiguous conclusions to be made. It has also been suggested that leaf nodule bacteria produce plant growth hormones, particularly cytokinins, which are necessary for the normal functioning of the plant (Fletcher and Rhodes-Roberts, 1976; reference is not an exact matchRodrigues-Pereira et al., 1972). Other authors claimed that leaf nodule bacteria can fix nitrogen, but this was contested by Van Hove (1976) and Lersten and Horner (1976). The lack of ability to fix nitrogen is also indicated by the absence of the nif HDK-like genes (Lambert et

749

Applications No applications of the genera Phyllobacterium or Ochrobactrum are yet known. However, the fact that Phyllobacterium is a predominant bacterium on the root surface of sugar beet plants, its capacity to “communicate” with plant tissues, and its non-pathogenic status make this bacterium an interesting new candidate for use in plant growth promotion or biological control of soil-borne diseases. Indeed, some of the sugar beet isolates exert a broad-spectrum antifungal activity against major phytopathogenic fungi.

Literature Cited Appelbaum, P. C., D. B. Campbell. 1980. Pancreatic abscess associated with Achromobacter group Vd biovar 1. J. Clin. Microbiol. 12:282–283. Barson, W. J., B. A. Cromer, M. J. Marcon. 1987. Puncture wound osteochondritis of the foot caused by CDC group Vd. J. Clin. Microbiol. 25:2014–2016. Chester, B., L. H. Cooper. 1979. Achromobacter species (CDC group Vd): morphological and biochemical characterization. J. Clin. Microbiol. 9:425–436. Clark, W. A., D. G. Hollis, R. E. Weaver, P. Riley. 1984. Identification of unusual pathogenic Gram-negative aerobic and facultatively anaerobic bacteria. Centers for Disease Control, Atlanta.

750

J. Swings et al.

Dees, S. B., C. W. Moss. 1978. Identification of Achromobacter species by cellular fatty acids and by production of keto acids. J. Clin. Microbiol. 8:61–66. De Jongh, P. 1938. On the symbiosis of Ardisia crispa (Thunb.). A. DC. Verh. Kon. Ned. Akad. Wetensch. Afd. Natuurk. Tweede Sect. II 37:1–74. De Ley, J., W. Mannheim, P. Segers, A. Lievens, M. Denijn, M. Vanhoucke, M. Gillis. 1987. Ribosomal ribonucleic acid cistron similarities and taxonomic neighborhood of Brucella and CDC Group Vd. Int. J. Syst. Bacteriol. 37:35–42. De Ley, J., P. Segers, K. Kersters, W. Mannheim, A. Lievens. 1986. Intra- and intergeneric similarities of the Bordetella ribosomal ribonucleic acid cistrons: proposal for a new family, Alcaligenaceae. Int. J. Syst. Bacteriol. 36:405–414. Fletcher, L. M., M. E. Rhodes-Roberts. 1976. The bacterial leaf nodule association in Psychotria. 99–118. D. W. Lovelock (ed.) Soc. appl. bacteriol. Technical series no 12. Academic Press, London. Gilardi, G. L. 1978. Identification of miscellaneous glucose non-fermenting Gram-negative bacteria. 45–65. G. L. Gilardi (ed.) Glucose non-fermenting gram-negative bacteria in clinical microbiology. CRC Press, Inc. Boca Raton, FL. Holmes, B., M. Popoff, M. Kiredjian, K. Kersters. 1988. Ochrobactrum anthropi gen. nov., sp. nov. from human clinical specimens and previously known as group Vd. Int. J. Syst. Bacteriol. 38:406–416. Horner, H. T., N. R. Lersten. 1972. Nomenclature of bacteria in leaf nodules of the families Myrsinaceae and Rubiaceae. Int. J. Syst. Bacteriol. 22:117–122. Kersters, K., J. De Ley. 1984. Genus III. Agrobacterium. 244– 254. N. R. Krieg and J. G. Holt (ed.) Bergey’s manual of systematic bacteriology, vol. 1. Williams & Wilkins, Baltimore. Knösel, D. 1962. Prü fung von Bakterien auf Fähigkeit zur Sternbildung. Zentralbl. Bakteriol. Parasitenk. Infektionskr. Hyg. Abt. 2 116:79–100. Knösel, D. 1984. The genus Phyllobacterium. 254–256. N. R. Krieg, and J. G. Holt (ed.) Bergey’s manual of systematic bacteriology, vol. 1. Williams & Wilkins, Baltimore.

CHAPTER 3.2.9 Lambert, B, H. Joos, S. Dierickx, R. Vantomme, J. Swings, K. Kersters, M. Van Montagu. 1990. The identification and plant interaction of a Phyllobacterium sp. a predominant rhizobacterium of young sugar beet plants. Appl. Environ. Microbiol. 56:1093–1102. Lersten, N. R., H. T. Horner, Jr. 1976. Bacterial leaf nodule symbiosis in angiosperms with emphasis on Rubiaceae and Myrsinaceae. Bot. Rev. 42:145–214. Miehe, H. 1911. Die Bakterienknoten an den Blatträndern der Ardisia crispa A. DC. Javanische Studien V. Abh. Math. Phys. Kl. Königl. Sächs. Gesellsch. Wiss. Leipzig, 32:399–431. Rodriguez-Pereira, A. S., P. J. W. Houwen, H. W. J. Deurenberg-Vos, B. F. Dey. 1972. Cytokinins and the bacterial symbiosis of Ardisia species. Zeitschr. Pflanzenphysiol. 68:170–177. Rubin, S. J., P. A. Granata, B. L. Wasilanskas. 1985. Glucosenonfermenting Gram-negative bacteria. 330–349. A. Balows, W. J. Hausler, Jr., and H. J. Shadomy (ed.) Manual of clinical microbiology, 4th ed. American Society for Microbiology. Washington, D.C. Stackebrandt, E., R. G. E. Murray, H. G. Trü per. 1988. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “purple bacteria and their relatives.” Int. J. Syst. Bacteriol. 38:321–325. Tatum, H. W., W. H. Ewing, R. E. Weaver. 1974. Miscellaneous Gram-negative bacteria. 270–294. E. H. Lennette, E. H. Spaulding, and J. P. Truant (ed.) Manual of clinical microbiology, 2nd ed. American Society for Microbiology. Washington, D.C. Van Hove, C. 1976. Bacterial leaf symbiosis and nitrogen fixation. 551–560. P. S. Nutman (ed.) Symbiotic nitrogen fixation in plants. Cambridge University Press. London, U.K. von Faber, F. C. 1912. Das erbliche Zusammenleben von Bakterien und tropischen Pflanzen. Jahrb. Wiss. Bot. 51:285–375. von Graevenitz, A. 1985. Ecology, clinical significance, and antimicrobial susceptibility of infrequently encountered glucose-nonfermenting Gram-negative rods. 181–232. G. L. Gilardi (ed.) Nonfermentative Gram-negative rods. Laboratory identification and clinical aspects. Marcel Dekker, Inc.. New York.

Prokaryotes (2006) 5:751–757 DOI: 10.1007/0-387-30745-1_34

CHAPTER 3.2.10 ehT

suneG

a i xreD

The Genus Derxia JAN HENDRIK BECKING

In the genus Derxia only a single species is recognized, D. gummosa. This species was originally described by Jensen et al. (1960) from an isolate obtained from Indian soil, but was later found to be widely distributed in soils of South America (Brazil), Southern Africa, Indonesia, and China, but not in temperate regions. Although Roy and Sen (1962) described a second species of Derxia (D. indica sp. nov.) from a sample of partially retted jute plant (Corchorus olitorius L.) from Uttar Pradesh, India, they did not present cultural and physiological data showing that their isolate was sufficiently different from Derxia gummosa to warrant the designation of a new species. The genus is named for H. G. Derx (1894–1953), a Dutch microbiologist. Using rRNA cistron similarity as a criterion for genetic relatedness, De Smedt et al. (1980) found that D. gummosa is quite different from species of the genera Azotobacter, Azomonas, and Beijerinckia. From hybridization studies with DNAs from three D. gummosa strains using the 14C-labeled rRNAs from a great variety of other organisms, it was quite obvious that the Derxia rRNA cistrons most closely resemble those of Pseudomonas acidovorans, P. solanacearum, Chromobacterium violaceum, Janthinobacterium lividum, and Alcaligenes faecalis. These taxa, together with a few others, constitute the third rRNA superfamily of De Ley (see The Proteobacteria: Ribosomal RNA Cistron Similarities and Bacterial Taxonomy in the second edition). Members of these taxa have in common that they consist of Gram-negative rods, usually 1.0–4.0 µm in length and 0.5–1.0 µm in diameter, do not possess resting stages, have a GC content of the DNA of 57–72 mol%, are chemoheterotrophic, exhibit respiratory (oxidative) metabolism, and occur in soil and water habitats. The third rRNA superfamily of De Ley (De Smedt et al., 1980) is identical with the beta subclass of the Proteobacteria, according to the phylogenetic taxon nomenclature of Stackebrandt et al. 1988. Based on numerical analysis of a large number of attributes, Thompson and Skerman (1979) showed fusion of their five Derxia strains tested with “Azotomonas” (“Azotomonas insolita” Stapp, 1940, sometimes termed “Pseudomonas insolita” [Stapp] Brisou, 1961) at group 1296 in their dendrograms of hierarchical interrelations (see also Krieg and Holt, 1984). This indicates that Derxia is not related to the genera Azotobacter or Azomonas and thus confirms the conclusions of De Smedt et al. (1980).

Habitats Derxia is primarily a soil organism occurring in tropical soils. There is no plausible explanation This chapter was taken unchanged from the second edition.

for why this organism is exclusively found in tropical environments and is apparently absent from temperate soils. It has not yet been recorded from aquatic habitats, although it is likely that it also occurs there. Such habitats in these regions have, however, not yet been thoroughly investigated using conditions selective for this organism.

Isolation The original isolate was obtained from a West Bengal (Adisaptagram, India) soil of pH 6.5 by the sieved-soil plate method (see “General Enrichment Procedures” in The Family Azotobacteraceae in the second edition) using plates with a nitrogen-free, mineral mannitol (2%) agar, incubated at room temperature (about 25°C). Yellowish colonies appeared after 5 days around some of the soil particles; these colonies gradually increased in size and finally assumed a rust-brown color. Isolation and further purification were performed by repeated transfer and plating on the same nitrogen-free mannitol agar. Campêlo and Döbereiner (1970) obtained this organism from Brazilian soils also with the sieved-soil plate method, utilizing a nitrogenfree, mineral starch (2%) medium. This medium was a modification of a medium used by Lipman (1903), supplemented with NaHCO3 (0.1 or 0.01%). Isolation Medium for Derxia gummosa (Campêlo and Döbereiner, 1970; Modified from Lipman, 1903) Distilled water Starch K2HPO4 KH2PO4 MgSO4 ·7H 2O CaCl2 NaHCO3 FeCl3 (10%, aqueous solution) Na2MoO4 ·2H 2O Bromothymol blue (0.5%, ethanol solution) Agar

1 liter 20.0 g 0.05 g 0.15 g 0.20 g 0.02 g 0.1 g 1 drop 0.002 g 5 ml 20.0 g

In a survey of 100 samples of Brazilian soil from the states of São Paulo, Rio de Janeiro,

752

J.H. Becking

CHAPTER 3.2.10

Pernambuco, and Para, Campêlo and Döbereiner (1970) found that 36% of the soils tested were positive for this organism. Soil humidity seems to favor the presence of Derxia, since 77% of the flooded soils, 36% of the humid soils, and only 13% of the drier soils tested contained this organism. In a comparative study with agar plates seeded with soil or with root pieces of various plants (mostly Gramineae) of the same locality, Derxia appeared more frequently on the plates inoculated with roots (33%) compared to those inoculated with soil (26%). D. gummosa was found to be present in soils ranging in pH from 4.5 to 6.5, but was observed to be most frequent in soils of pH 5.1–5.5. This species was isolated by the author from various soils of Indonesia, China, Brazil, tropical Africa, and South Africa with the sieved-soil plate method using nitrogen-free, mineral, glucose (1–2%) agar (J. H. Becking, unpublished observations). Derxia strains are relatively acid tolerant and were therefore also isolated by the author from cultures designed to enrich for Beijerinckia strains using a liquid, acidic, nitrogen-free, glucose medium. This liquid medium has the following composition: Enrichment Medium for Derxia gummosa (J. H. Becking, unpublished observations) Distilled water Glucose KH2PO4 MgSO4 ·7H 2O

1 liter 20.0 g 1.0 g 0.5 g

Adjust to pH 5.0.

The medium is dispensed into petri dishes so as to form a thin layer 2–3 mm deep in order to maintain aerobic conditions. The liquid medium is inoculated with soil particles and incubation is at 25° or 30°C. In the sieved-soil plate method mentioned for the isolation, soil particles are evenly distributed over the surface of nitrogen-free mineral agar containing mannitol (Jensen et al., 1960), glucose (J. H. Becking, unpublished observations), or starch (Campêlo and Döbereiner, 1970). It should be noted that mannitol is not utilized by all Derxia gummosa strains. Of the six strains tested by Thompson and Skerman (1979), three did not use mannitol. Moreover, some strains, including the type strain, produced only very scant growth on starch. All strains did, however, utilize glucose as sole source of carbon. Therefore for general practice of the sievedsoil plate method, the following glucose mineral agar medium is recommended: Nitrogen-free, Mineral Glucose Agar for the Isolation of Derxia gummosa with the Sieved-Soil Plate Method (g/liter of distilled water):

Glucose K2HPO4 KH2PO4 MgSO4 ·7H 2O FeCl3 ·6H 2O Na2MoO4 ·2H 2O CaCl2 Agar

20.0 0.8 0.2 0.5 0.025 (or 0.05) 0.005 0.05 15.0

Adjust to pH 6.9.

On the agar plates seeded with the soil particles, yellowish colonies develop around the soil particles. These colonies eventually become larger and acquire a rust-brown color. Isolates should be further purified by repeated streaking on new agar plates. For agar plates obtained from liquid enrichment cultures, one should look for slimy, yellowish colonies with a rather plicated surface (see Fig. 5).

Cultivation For routine maintenance, the following nitrogenfree, mineral glucose medium can be used (g/liter of distilled water): Nitrogen-free, Mineral Glucose Agar Medium Glucose K2HPO4 MgSO4 ·7H 2O NaCl FeSO4 ·7H 2O CaCl2 or CaCO3 Na2MoO4 ·2H 2O Agar

10.0 (or 20.0) 0.5 0.25 (or 0.2) 0.25 0.1 0.1 0.005 15.0

Adjust to pH 6.9.

Jensen et al. (1960) used a nitrogen-free, mineral mannitol agar of the following composition (g/liter of distilled water): Nitrogen-free, Mineral Mannitol Agar Medium Mannitol K2HPO4 MgSO4 ·7H 2O CaCl2 CaCO3 Na2WO4 ·2H 2O FeCl3 and Na2MoO4 ·2H 2O Agar

10.0 0.5 0.2 0.1 5.0 0.0005 trace 15.0.

The medium recommended by Campêlo and Döbereiner (1970) contained starch as carbon source (g/liter of distilled water): Nitrogen-free, Mineral Starch Agar Medium Starch K2HPO4 KH2PO4 MgSO4 ·7H 2O CaCl2 NaHCO3 FeCl3 solution (10%, aqueous solution)

20.0 g 0.05 g 0.15 g 0.2 g 0.02 g 0.1 g 1 drop

CHAPTER 3.2.10 Na2MoO4 ·2H 2O Bromthymol blue solution (0.5%, ethanol solution) Agar

The Genus Derxia 0.002 g 5.0 ml 20.0.

As already stated before, not all Derxia gummosa strains can utilize mannitol as sole source of carbon, while with starch as the carbon source, some strains only produce scant growth.

753

of Derxia acquire on aging a typical dark rustbrown or mahogany-brown color. In addition, in contrast to Azotobacter, Azomonas and Beijerinckia, Derxia cells are catalase negative. In contrast to the spiral-shaped or vibrioid cells of Azospirillum (see The Genera Azospir-

Preservation of Cultures The maintenance procedures of Derxia gummosa are the same as those described for the genus Beijerinckia (see The Genus Beijerinckia in Volume 5).

Identification Derxia can be distinguished from other genera of N2-fixing bacteria by its very slimy (gummy) growth, both on agar plates and in liquid media, combined with the very pleomorphic appearance of the cells (Figs. 1 to 2, 3, 4) depending on age and type of medium. On agar media Azotobacter and Azomonas species never produce such slimy colonies as those of Derxia (Fig. 5). Confusion with Beijerinckia is unlikely, since Beijerinckia cells usually show very characteristic cells with only two polar lipoid bodies, whereas Derxia cells contain numerous lipoid bodies. Moreover, in contrast to Beijerinckia colonies, the colonies

Fig. 2. Three-week-old cells of Derxia gummosa on nitrogenfree glucose agar, showing shrinkage of the cells. Phase contrast microscopy. Bar = 10 µm.

Fig. 1. Seven-day-old cells of Derxia gummosa on nitrogenfree agar containing 2% glucose. Phase contrast microscopy. Bar = 10 µm.

Fig. 3. Ten-day-old cells of Derxia gummosa on peptone agar with 2% glucose. Phase contrast microscopy. Bar = 10 µm.

754

J.H. Becking

CHAPTER 3.2.10

a

b

Fig. 5. Colony of Derxia gummosa on nitrogen-free glucose agar with calcium carbonate showing the coarse and wrinkled colony surface. Bar = 1 cm.

Fig. 4a. Three-month-old cells of Derxia gummosa on nitrogen-free glucose agar. The cells show shrinkage and are enclosed by a thick slime envelope. Phase contrast microscopy. Bar = 10 µm. Fig. 4b. More pronounced shrinkage of cells of a four-month-old Derxia gummosa culture on nitrogen-free glucose agar. Note the thick slime envelopes enclosing the rows of cells. Bar = 10 µm.

illum and Herbaspirillum in the second edition), Derxia cells are mainly straight rods and the lipoid bodies are more numerous. Derxia colonies are far more slimy than those of Azospirillum, and the mahogany-brown color they eventually acquire is in contrast to the pink or whitish color of Azospirillum colonies. Moreover, there are some minor physiological differences. Derxia strains can exhibit nitrogendependent growth on glucose, whereas only some strains of Azospirillum (i.e., only those belonging to the species A. lipoferum) can do so. On the other hand, Derxia strains generally do not grow well on malate, whereas all strains of Azospirillum can grow well on this carbon source. Moreover, Derxia strains are catalase negative, whereas Azospirillum strains give reactions that range from strong to undetectable.

Unlike Azotobacter, Azomonas, and Beijerinckia species, Derxia species can grow as a facultative hydrogen autotroph. In this ability, they show affinities to various other facultative hydrogen autotrophs that can also use H2 to provide energy and reducing power for growth and CO2 fixation, i.e., various Pseudomonas and Alcaligenes species (“Hydrogenomonas” species) (Buchanan and Gibbons, 1974), Xanthobacter species (Wiegel et al., 1978; Wiegel and Schlegel, 1984; Malik and Claus, 1979), nitrate-reducing Paracoccus species (Rittenberg, 1969), Bradyrhizobium japonicum (Hanus et al., 1979), and Azospirillum lipoferum (Malik and Schlegel, 1981; Sampaio et al., 1981) (see also The Mesophilic Hydrogen-Oxidizing (Knallgas) Bacteria in the second edition). For further details on facultative hydrogen autotrophy of Derxia, see section “Physiological and Biochemical Properties” in this Chapter.

Species Description Derxia gummosa Jensen, Petersen, De and Bhattacharya, 1960. Cells are Gram-negative, rod-shaped with rounded ends, 3.0–6.0 µm in length, and 1.0–

CHAPTER 3.2.10

1.2 µm wide, occurring singly or in short chains. Cells are rather pleomorphic, depending on age and the medium. In aging cultures, cells often remain together, forming long filaments of sometimes locally swollen or distorted cells. Some cells may assume enormous sizes (up to 30 µm). Young cells have a homogeneous cytoplasm; older cells typically show large refractile bodies throughout the whole cell. Resting stages are not known. Cells are motile by a short polar flagellum; motile cells are numerous in liquid glucose media containing combined nitrogen, but are rare on nitrogen-deficient solid media. The organism is aerobic, having a strictly respiratory type of metabolism with oxygen as the terminal electron acceptor. Molecular nitrogen is fixed under aerobic conditions and also under decreased oxygen pressures (microaerophilic conditions). Optimum temperature is 25–35°C; growth is slow at 15°C, feeble at 40°C; no growth occurs at 50°C. Growth occurs from pH 5.5 to about 9.0; no growth occurs at pH 4.4. Liquid cultures turn into a gelatinous mass, but growth near the surface is more luxuriant and forms a thick, tough pellicle. Colonies on agar media are at first slimy and semitransparent, later massive and opaque, highly raised with a wrinkled surface. Older colonies develop a rust-brown to dark mahoganybrown color. They are catalase negative. A wide range of sugars, alcohols, and organic acids are oxidized, mostly to CO2, but a small amount of acid, probably acetic acid, is produced during growth in an alkaline medium. They can grow as a facultative hydrogen autotroph. The GC content of the DNA in three strains of Derxia gummosa ranges from 69.2–72.6 mol% (Tm) (De Smedt et al., 1980). In the type strain, it is 70.4 ± 1.7 mol% (Tm) (De Ley and Park, 1966). The appearance of cells from young cultures is depicted in Fig. 1. Older cells on sugar-rich media contain large refractile bodies. Especially on glucose-peptone agar, very elongated cells are produced containing many refractile bodies (Fig. 3). The refractile material is probably poly-βhydroxybutyrate, since it stains with Sudan III and Sudan black, but some vacuoles which do not stain may also be involved. On nitrogen-free glucose agar, older cells undergo shrinkage (Fig. 2) and are finally enclosed by a slime envelope (Fig. 4). Motile cells may become numerous in liquid glucose media containing ammonia or glutamate as the nitrogen source. The cells usually have a single polar flagellum, but some may have a flagellum at each pole. According to Thompson and Skerman (1979), the single polar flagellum is less than a full wave and is less than 3 µm in length. Growth in liquid media usually starts as a ring at the glass-liquid interface and develops into a thick, wrinkled, tough pellicle. Shallow layers of

The Genus Derxia

755

medium change into a firm gelatinous mass after a couple of weeks. The color gradually becomes a dark red-brown. Growth on nitrogen-deficient agar media begins as thin, whitish, or semitransparent scattered colonies. Later, more massive, highly raised, or dome-shaped colonies emerge which rapidly assume a diameter of 1 cm or more (giant colonies) (Fig. 5). These colonies are very reminiscent of those of Beijerinckia species. Colonies are at first whitish or dull yellow with a smooth surface, but the surface soon becomes coarse and wrinkled, and the color deepens to a dark mahogany brown. The slime of these colonies is very tenacious and gumlike, but in the other developmental stages, it is softer and smeary.

Physiological and Biochemical Properties In the original description, Jensen et al. 1960 mentioned the appearance of Derxia gummosa on nitrogen-free, mineral glucose agar as a few “massive” colonies among many “thin,” whitish colonies. The type of colony used as inoculum affected neither the relative numbers nor the distribution of the two colony types. They reported further that the “massive” colonies fixed nitrogen, whereas the “thin” colonies did not. Tchan and Jensen (1963) observed that the formation of “massive” colonies was stimulated by the addition of small quantities of combined nitrogen to the medium and that dinitrogen fixation could not be initiated in a medium completely free of combined nitrogen. Hill and Postgate (1969) found that, with a small quantity of combined nitrogen, batch cultures of Derxia gummosa in otherwise nitrogenfree media could be established with agitation in air. However, this supplement was unnecessary if the cultures were left stagnant or if the atmospheric oxygen concentration was lowered to 0.1 atm or less. Once established, continuous cultures could be maintained in air, provided vigorous stirring was avoided. Hill (1971) demonstrated in more detail that D. gummosa is very sensitive to oxygen and that the presence of oxygen markedly affects its colony morphology. At an oxygen concentration of less than 0.2 atm, only the “massive”-colony type is produced on nitrogen-free, mineral agar media; the “thin”colony type which predominates in air was absent. Only the “massive” colonies reduced acetylene. Apparently, the copious production of a viscous and tenacious slime inhibits the penetration of oxygen and therefore stimulates nitrogen fixation. The colonial dimorphism on agar probably arises because only when the local oxygen concentrations on the agar surface are

756

J.H. Becking

depressed can dinitrogen fixation and subsequent growth to the “massive” colony type take place. The efficiency of nitrogen fixation by D. gummosa varies between 9 and 25 mg N/g of glucose consumed, but in most strains it is distinctly lower than in Azotobacter or Beijerinckia. Growth on nitrogen-deficient media or under nitrogen-fixing conditions is stimulated by small amounts of combined nitrogen (nitrate, ammonia), particularly at the start (Tchan and Jensen, 1963). There is no requirement for amino acids under nitrogen-fixing conditions. However, small amounts of yeast extract are stimulatory for growth under nitrogen-fixing conditions (J. H. Becking, unpublished observations), and although there is no apparent requirement for vitamins or growth factors, it is possible that small amounts of biotin are stimulatory for growth of some strains (J. H. Becking, unpublished observations). As sole source of carbon, growth is good to excellent on glucose, fructose, ethanol, glycerol, mannitol (only utilized by some strains), and sorbitol; growth on lactate is scant. No growth or only a trace of growth occurs on lactose, galactose, maltose, sucrose, formate, acetate, propionate, pyruvate, succinate, malate, fumarate, dulcitol, and sometimes starch. Butyrate, citrate, benzoate, and xylose suppress growth. Growth on methane or methanol as the sole carbon source has also been demonstrated for D. gummosa (Sampaio et al., 1981). Growth with combined nitrogen sources is much faster than with molecular nitrogen and is completely uniform, in contrast to the uneven growth on nitrogen-free agar. Colonies change from a pale yellow through rust-brown to almost black (darkest if nitrate is present), and sometimes a light brown, water-soluble pigment is produced. Growth with glutamic acid, ammonium acetate, alanine, sodium nitrate, and urea decreases from abundant to good in approximately the same sequence. Aspartic acid, asparagine, and peptone give a much slower growth, which is uneven and mostly confined to scattered colonies. Glycine seems to be toxic. Nitrate is not reduced to nitrite or N2 in a glucose-nitrate medium. Indole is not produced from tryptophan. Trace elements, particularly molybdenum, are required for dinitrogen fixation, but vanadium cannot replace molybdenum (Jensen et al., 1960). Thus, Derxia apparently does not have an alternative vanadium-activated nitrogenase. D. gummosa has been shown to grow as a hydrogen autotroph in an atmosphere containing H2 + CO2 + O2, with either N2 or NH4+ as the nitrogen source (Pedrosa et al., 1980). Indeed, it appears to grow nearly as well autotrophically as

CHAPTER 3.2.10

it does heterotrophically. Ribulose-1,5-bisphosphate carboxylase activity, which mediates CO2 fixation, occurs in autotrophically-grown cells but not in cells grown heterotrophically. A medium for testing the autotrophy has the following composition (g/liter of distilled water): Medium for Testing Autotrophy KH2PO4 K2HPO4 MgSO4 ·7H 2O NaCl CaCl2 ·2H 2O FeSO4 ·7H 2O Trace element solution (see below) Biotin Agar

1.2 0.8 0.2 0.2 0.02 0.002 2.0 ml 10 µg 15.0

Trace element solution (g/liter) Na2MoO4 ·2H 2O MnSO4 ·H 2O H3BO3 CuSO4 ·5H 2O ZnSO4 ·7H 2O

1.0 1.75 1.4 0.04 0.12

This medium was employed by Pedrosa et al. (1980) in their studies on the autotrophy of Derxia. In these studies, a very low concentration of potassium malate (0.1 g/liter) was sometimes added to the medium. It should be noted that most strains of Derxia (including the type strain) give only scant growth with malate. Thompson and Skerman (1979) reported no growth on DLmalate in all of the six strains that they tested.

Applications No studies have yet been done on the occurrence and significance of this organism in the rhizosphere of crop plants. However, Campêlo and Döbereiner (1970) found Derxia more frequently on agar plates inoculated with roots (33% positive for Derxia) compared to those inoculated with soil (26%).

Literature Cited Brisou, B. 1961. Etudes de quelques Pseudomonas chromogènes isolés àDiego-Suarez. Bull. Soc. Path. Exot. 54:746–755. Buchanan, R. E., N. E. Gibbons (ed.). 1974. Bergey’s manual of determinative bacteriology. 8th ed. Williams and Wilkins. Baltimore. Campêlo, A. B., Döbereiner, J. A. 1970. Ocorrência de Derxia sp. em solos de alguns Estados Brasileiros. Pesquisa Agropecuá ria Brasileira 5:327–332. De Ley, J., Park, I. W. 1966. Molecular biological taxonomy of some free-living nitrogen-fixing bacteria. Antonie van Leeuwenhoek, Journal of Microbiology and Serology 32:6–16. De Smedt, J., Bauwens, M., Tytgat, R., De Ley, J. 1980. Intraand intergeneric similarities of ribosomal ribonucleic

CHAPTER 3.2.10 acid cistrons of free-living, nitrogen-fixing bacteria. Int. J. Syst. Bacteriol. 30:106–122. Hanus, F. J., Maier, R. J., Evans, H. J. 1979. Autotrophic growth of H2-uptake-positive strains of Rhizobium japonicum in an atmosphere supplied with hydrogen gas. Proc. Natn. Acad. Sci. USA. 76:1788–1792. Hill, S. 1971. Influence of oxygen concentration on the colony type of Derxia gummosa grown on nitrogen-free media. J. Gen. Microbiol. 67:77–83. Hill, S., Postgate, J. R. 1969. Failure of putative nitrogenfixing bacteria to fix nitrogen. J. Gen. Microbiol. 58:277– 285. Jensen, H. L., Petersen, E. J., De, P. K., Bhattacharya, R. 1960. A new nitrogen-fixing bacterium: Derxia gummosa nov. gen. nov. spec. Arch. Microbiol. 36:182– 195. Krieg, N. R., Holt, J. G. (ed.). 1984. Bergey’s manual of systematic bacteriology, vol. 1. Williams and Wilkins. Baltimore. Lipman, J. G. 1903. Experiments on the transformation and fixation of nitrogen by bacteria. 217–285. 16th Annual Report over 1903 of the New Jersey State Agricultural Experiment Station. USA. Malik, K. A., Claus, D. 1979. Xanthobacter flavus, a new species of nitrogen-fixing hydrogen bacteria. Int. J. Syst. Bacteriol. 29:283–287. Malik, K. A., Schlegel, H. G. 1981. Chemolithotropic growth of bacteria able to grow under N2-fixing conditions. FEMS Microbiol. Lett. 11:63–67. Pedrosa, F. O., Döbereiner, J., Yates, M. G. 1980. Hydrogendependent growth and autotrophic carbon dioxide fixation in Derxia. J. Gen. Microbiol. 119:547–551.

The Genus Derxia

757

Rittenberg, S. C. 1969. The roles of exogenous organic matter in the physiology of chemolithotrophic bacteria. Adv. Microbiol. Physiol. 3:159–196. Roy, A. B., Sen, S. 1962. A new species of Derxia. Nature London, 194:604–605. Sampaio, M.-J. A. M., da Silva, E. M. R., Döbereiner, J., Yates, M. G., Pedrosa, F. O. 1981. Autotrophy and methylotrophy in Derxia gummosa, Azospirillum brasilense and A. lipoferum. 447. Gibson, A. H., and Newton, W. E. (ed.) Current perspectives in nitrogen fixation, Proc. 4th Int. Symp. on Nitrogen Fixation, Canberra, Australia, Dec. 1–5, 1980. Elsevier/North-Holland Biomedical Press. Amsterdam. Stackebrandt, E., Murray, R. G. E., Trü per, H. G. 1988. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “purple bacteria and their relatives.” Int. J. Syst. Bacteriol. 38:321–325. Stapp, C. 1940. Azotomonas insolita, ein neuer aerober stickstoffbindender Mikroorganismus. Zentralblatt fü r Bakteriologie, Parasitenkunde, Infektionskrankheiten, und Hygiene, Abt. II 102:142–150. Tchan, Y. T., Jensen, H. L. 1963. Studies of nitrogen fixing bacteria. Proc. Linn. Soc. New South Wales 88:379–385. Thompson, J. P., Skerman, V. B. D. 1979. Azotobacteraceae: The taxonomy and ecology of the aerobic nitrogenfixing bacteria. Academic Press. London. Wiegel, J., Schlegel, H. G. 1984. Genus Xanthobacter. 325– 333. Bergey’s manual of systematic bacteriology. Williams and Wilkins. Baltimore. Wiegel, J., Wilke, D., Baumgarten, J., Opitz, R., Schlegel, H. 1978. Transfer of the nitrogen-fixing hydrogen bacterium Corynebacterium autotrophicum Baumgarten et al. to Xanthobacter gen. nov. Int. J. Syst. Bacteriol. 28:573–581.

Prokaryotes (2006) 5:758–777 DOI: 10.1007/0-387-30745-1_35

CHAPTER 3.2.11 ehT

areneG

x i rhtotpeL

dna

su l i toreahpS

The Genera Leptothrix and Sphaerotilus STEFAN SPRING

Introduction Representatives of the genera Leptothrix and Sphaerotilus were among the first microorganisms to be recognized in the environment and described in detail by scientists. The type species of the genus Leptothrix, L. ochracea, was already observed in the late eighteenth century and described by Roth (1797) under the synonym “Conferva ochracea.” Later, Kü tzing (1843) proposed to place this species within the genus Leptothrix. Ten years earlier, the same author had published a description of the species Sphaerotilus natans (Kü tzing, 1833), which is today still known under this name. These early publications were probably evoked by the observation of ocherous deposits (clearly visible to the naked eye) in ponds or slowly running water. A microscopic examination of these suspicious aggregates led then to the discovery of filamentous microorganisms, which were obviously responsible for the deposition of iron or ferromanganese oxides in a slimy matrix, resulting in the typical color of the formed amorphous flocs or surface films. Further investigations of the SphaerotilusLeptothrix group were induced by their potential economic importance. The massive growth of these microorganisms at industrial sites can lead to technological problems like clogging of water distribution systems or the bulking of activated sludge (Dondero, 1975). More recently, the significance of Leptothrix species in the corrosion of steel could be demonstrated (Olesen et al., 2000; Rao et al., 2000). On the other hand, a potential application of these microorganisms in the biological clearance of heavy metals from contaminated water supplies is being discussed (Nelson et al., 1999; Solisio et al., 2000). Both genera, Leptothrix and Sphaerotilus, have been always considered as closely related because of the conformity of some suspicious morphologic traits (Mulder and Van Veen, 1963). Later, their close relationship was confirmed by phylogenetic and phenotypic data, thereby justifying their treatment as a group. Characteristic traits, which distinguish members of this group

from other phylogenetically related species, are the capability to form tubular sheaths and the precipitation of copious amounts of oxidized iron or manganese.

Phylogeny and Taxonomy Phylogeny and Related Genera Members of the genera Leptothrix and Sphaerotilus comprise a phylogenetically coherent cluster within the β1-subgroup of the Proteobacteria. The similarity values among 16S rRNA gene sequences representing strains of the Sphaerotilus-Leptothrix group are in the range between 96.3 and 99.8% (Pellegrin et al., 1999). It has to be noted, however, that to date no 16S rRNA sequence of L. ochracea (the type species of the genus Leptothrix) is available. In addition, the physiology of this species is largely unknown, because it could not be isolated in pure culture. Hence, it cannot be excluded that the type species of Leptothrix is phylogenetically only distantly related to the other species of this genus. Representatives of the genera Aquabacterium, Ideonella, Rubrivivax, Roseateles and two misclassified species, [Alcaligenes] latus and [Pseudomonas] saccharophila, are phylogenetically closely related to the SphaerotilusLeptothrix group. Together they form the Rubrivivax line of descent, which is tightly associated with the Comamonadaceae (Wen et al., 1999). The relative branching order of most species within the Rubrivivax group is difficult to determine, mainly because of the restricted number of varying positions available for the estimation of phylogenetic distance values. Depending on the method and database used for tree reconstruction, it may happen that representatives from other genera get intermixed with species from the Sphaerotilus-Leptothrix group. Therefore, it is difficult to decide if the capability of sheath-formation within this group evolved from a common ancestor, i.e., if it is a monophyletic trait. Despite the close phylogenetic relationship of members of the Rubrivivax branch, a differentiation is easily possible based on phenotypic

CHAPTER 3.2.11

The Genera Leptothrix and Sphaerotilus

759

Rubrivivax gelatinosus (M60682)

Ideonella dechloratans (X72724)

Aquabacterium parvum (AF035052) Aquabacterium commune (AF035054) Aquabacterium citratiphilum (AF035050)

[Pseudomonas] sacchorophila (AB021407) Leptothrix discophora (L33975) Leptothrix mobilis (X97071) Leptothrix cholodnii (X97070)

Roseateles depolymerans (AB003623) [Alcaligenes] latus (D88007)

Sphaerotilus natans (Z18534)

5%

Fig. 1. Phylogenetic tree showing the position of members of the Sphaerotilus-Leptothrix group among representatives of the Rubrivivax-branch of the β-Proteobacteria. Sheath-forming species are labeled in red, photoautotrophic species in green, and facultatively chemolithoautotrophic species in blue. The GenBank/EMBL accession number for each sequence is shown in parentheses. With the exception of Sphaerotilus natans (DSM 565) and Leptothrix cholodnii (CCM 1827), sequences of type strains were used. This tree was reconstructed using the ARB program package (Ludwig and Strunk, 1997). It is derived from a distance matrix on a selection of 16S rRNA sequences using the neighbor-joining method of Saitou and Nei (1987). Phylogenetic distances were calculated as described by Felsenstein (1982). The sequence of Escherichia coli was used as an outgroup (not shown). The bar indicates 5% estimated sequence divergence.

characteristics. It is noteworthy that the genera in this tight assemblage represent three basically different metabolic types, namely photoautotrophs (Rubrivivax), facultative chemolithoautotrophs ([Alcaligenes] latus and [Pseudomonas] saccharophila) and respiratory chemoorganotrophs (e.g., Ideonella and Sphaerotilus). In Figure 1, a representative phylogenetic tree based on 16S rRNA gene sequence data has been selected which adequately reflects the phenotypic classification within this group. Important phenotypic characteristics useful for distinguishing members of the genera Leptothrix and Sphaerotilus from other species belonging to the Rubrivivax-lineage are listed in Table 1.

Phenotypic Characteristics Common Traits Some important phenotypic traits are shared by all members of the genera Leptothrix and Sphaerotilus: Cells are straight rods, Gram-negative and motile by flagella. Growth is filamentous under natural conditions, owing to the formation of tubular sheaths that surround single and linear chains of cells. Sheaths are produced by excretion of fibrillar polymeric substances that are crosslinked to form a mesh-like fabric closely fitting to the cells. In contrast to slimes or capsules, this matrix is not in intimate contact with the cells. In aquatic habitats rich in iron and manganese, sheaths get incrusted with ferric hydroxide (Sphaerotilus) or ferromanganese oxides (Leptothrix). Without these incrustations, they appear as very thin,

almost transparent structures and are therefore not easily recognized by phase contrast microscopy. Occasionally, cells move out of the sheath or lyse and leave behind an empty space or gap within the filament, thereby facilitating the observation of the sheath. Treatment of slide preparations with ethanol may improve the visibility of sheaths under phase contrast. Further conspicuous morphological features of members of this group include the formation of slime, holdfasts, and false branching of filaments. These traits are frequently recognized in the genus Sphaerotilus, but are less common among representatives of the genus Leptothrix. The sheaths of S. natans and several Leptothrix species are surrounded by a slime layer that may be involved in the accumulation of iron hydroxide and in the oxidation of Mn(II). Holdfasts are sticky, disc-like evaginations or blebs formed at one cell pole opposite to the flagellum by which organisms attach to walls of containers, submerged plants, stones and other surfaces. Holdfast formation can be easily observed in pure cultures of Sphaerotilus natans and Leptothrix lopholea, but is absent or variable in other cultured Leptothrix species. False branching is characteristic for Sphaerotilus natans and several Leptothrix strains. It develops if a cell attaches to an existing filament and forms a new filament, thereby forming a branch. Metabolism is strictly aerobic, respiratory and chemoorganoheterotrophic. The possibility of an autotrophic or mixotrophic growth of these bacteria with ferrous iron as electron acceptor has been discussed for many years, but up to now no

69–71

Soil

69 River water

66

NR

−

−

−

+

−

Several polar

1

Roseateles

Mud

70–72

NR

+

+

+

+

−

One polar

1

Rubrivivax

Activated sludge

68

NR

+

NR

−

−

−

Several, polar or subpolar

1

Ideonella

Drinking water

Freshwater, sediment

68–71

+

− 65–66

−

NR

−

−

+

One polar; subpolar tuft

5

Leptothrixa

+

NR

−

−

−

One polar

3

Aquabacterium

Freshwater, activated sludge

69

−

−

NR

−

−

+

Subpolar tuft

1

Sphaerotilus

a

Symbols: +, present in all species; −, absent in all species; and NR, not reported. Physiological data for Leptothrix lopholea and L. ochracea are not available. From Kersters and De Ley (1984); Palleroni (1984); Mulder and Deinema (1992); Malmqvist et al. (1994); Spring et al. (1996); Kalmbach et al. (1999); and Suyama et al. (1999).

Isolation source

G + C content (mol%) Mud

−

−

Anaerobic growth NR

+

+

Autotrophic growth with H2

NR

−

−

Photoautotrophic growth

Oxidation of Mn

−

−

Carotenoid pigments

2+

−

One polar

1

[Pseudomonas] saccharophila

−

Peritrichous

1

[Alcaligenes] latus

Formation of sheaths

Flagellation

Number of species

Characteristic

Table 1. Differential characteristics of genera and misclassified species belonging to the Rubrivivax group of the β-Proteobacteria.

760 S. Spring CHAPTER 3.2.11

CHAPTER 3.2.11

definite evidence could be presented which would support this assumption. Representatives of both genera, Leptothrix and Sphaerotilus, require vitamin B12 as an essential growth factor in mineral media. A number of Leptothrix strains have been found to require in addition thiamine and biotin as growth factors (Rouf and Stokes, 1964). In broth culture, a flocculent growth is typical. Poly-β-hydroxybutyrate is stored as reserve material. The major quinone type detected is ubiquinone Q8 (>90% of total quinones). The four major components of the cellular fatty acids are cis-9 hexadecanoic acid (cis-9 16:1), hexadecanoic acid (16:0), cis-9,11 octadecanoic acid (cis-9,11 18:1) and decanoic acid (12:0). In all strains tested, the hydroxylated fatty acid 3-hydroxydecanoic acid (3-OH 10:0) can be detected, however in some cases, only in low amounts (Kämpfer, 1998). The G +C content of DNA is similar in both genera and ranges between 68 and 71 mol%. In general, chemotaxonomic features characteristic for the Sphaerotilus-Leptothrix group are very similar in other organisms belonging to the Rubrivivax group and Comamonadaceae, so that a clear differentiation of species or even genera within this phylogenetic group is hardly possible based solely on chemotaxonomic traits. Distinguishing Traits Albeit closely related, several phenotypic characteristics allow the differentiation of the genera Leptothrix and Sphaerotilus. The ability to oxidize soluble manganese (Mn2+) compounds to solid manganic (Mn4+) oxides is restricted to Leptothrix species. This trait has been originally used for differentiation because it can be easily observed in Mn2+-containing media or habitats. Further distinguishing traits, however, were only revealed by a detailed taxonomic study of cultured strains of both genera. They include the storage of polysaccharides as reserve material only by Sphaerotilus strains, size of cells, structure of the sheath surface (Leptothrix sheaths show a rough surface when viewed by electron microscopy, in contrast to the sheaths of Sphaerotilus strains which have a smooth surface), utilization of carbon sources, and the pronounced response of S. natans to an increase of organic nutrient concentration in contrast to a poor response of most Leptothrix species. A summary of phenotypic characteristics useful for the classification of both genera is presented in Table 2.

Problems in the Taxonomy of Members of the Sphaerotilus-Leptothrix Group The Proposal of Pringsheim Members belonging to the Sphaerotilus-Leptothrix group share

The Genera Leptothrix and Sphaerotilus

761

Table 2. Main morphological and physiological characteristics of the genera Leptothrix and Sphaerotilus. Characteristic Cell dimensions Width (µm) Length (µm) Flagella Monotrichous, polar Polytrichous, subpolar Structure of sheath surfaceb Reserve material Poly-β-hydroxybutyrate Polysaccharide Major fatty acidsc

Hydroxylated fatty acid Major quinone type Oxidation of Mn2+ Growth stimulation by increase of nutrient concentration Need for vitamin B12 Carbon sources used for growthe l-Alanine l-Asparagine l-Aspartate Butyrate d-Fructose d-Glucose d-Gluconate l-Ornithine

Leptothrixa

Sphaerotilus

0.6–1.5 1.5–14

1.2–2.5 1–10

+ + Rough

− + Smooth

+ − cis-9 16:1, 16:0, cis-9,11 18:1 3-OH 10:0 Q-8 + −/+d

+ + cis-9 16:1, 16:0, cis-9,11 18:1 3-OH 10:0 Q-8 − +

+

+

− − − − D D − −

+ + + + + + + +

Symbols: +, 90% or more strains are positive; −, 90% or more strains are negative; D, 11–89% of the strains are positive. a Results are based on strains available from culture collections. b Electron microscopic observation of unstained preparations. c Number of carbon atoms:number of double bonds. d Most freshly isolated strains show no pronounced response, with the exception of Leptothrix cholodnii strains. e Utilization of carbon sources was tested in GMBN medium (Richard et al., 1985; Kämpfer, 1998). From Van Veen et al. (1978); Spring et al. (1996); and Kämpfer (1998).

some important phenotypic characteristics which are easily recognized. On the other hand, distinguishing traits are sometimes difficult to determine and require the investigation of defined pure cultures. Certain strains of this group show a considerable morphological variability, depending on the respective environmental conditions, leading to the effect that virtually identical or similar strains were described under different species names. These circumstances probably led Pringsheim (Pringsheim, 1949a; Pringsheim, 1949b) to believe that all strains of this group should be placed in the genus Sphaerotilus. He also concluded that L. ochracea would be a morphological variant of S. natans

762

S. Spring

because both species lack visible deposits of manganese oxides on their sheaths. The investigations which led to these conclusions were however inadequate because they were based on only a few partly described strains or on the observation of uncultured bacteria in natural environments. Several studies in the following years clearly revealed the mistakes in Pringsheim’s nomenclature. Nevertheless, his assumptions produced a lot of confusion in the taxonomic literature on the SphaerotilusLeptothrix group between the 1950s and 70s. In that time, for instance, a variety of manganeseoxidizing strains, which probably belong to different Leptothrix species, were described under the name Sphaerotilus discophorus (e.g., Rouf and Stokes, 1964). Instability of Phenotypic Characteristics Several difficulties regarding the taxonomy of this group are due to the loss of important phenotypic traits upon cultivation (Van Veen et al., 1978). The reversible or nonreversible loss of physiological and morphological characteristics of strains in pure culture caused divergent descriptions of species, which were originally described on the basis of observations in the natural environment (e.g., L. discophora, see below). The ability to form sheaths has been irreversibly lost by several strains of Sphaerotilus natans and by most Leptothrix strains available from public culture collections, with the exception of L. cholodnii (formerly “L. discophora”) SP-6 (= LMG 8142). Gaudy and Wolfe (1961) reported that sheath formation in Spaerotilus natans is affected by the composition of the culture medium and inhibited at high concentrations of peptone. The colony morphology of freshly isolated strains corresponds to the ability of sheath formation and can be smooth (sheathless cells) or rough and filamentous (sheathforming cells). Sometimes even various colony morphologies of the same strain can be observed on one agar plate. The manganese-oxidizing activity in Leptothrix strains is independent from the formation of sheaths and more stable in most strains, but loss of this trait has also been reported. A report on the disc- or holdfast-formation of an uncultured sheathed bacterium, which was tentatively identified as L. discophora (Carlile and Dudeney, 2001), illustrates the taxonomic problems within this group. Interestingly, the formation of basal discs or holdfasts at one end of the filament (discophora means disc-bearing) was mentioned in the original description of this species by Schwers (1912), which was based on photomicrographs of cells in natural environments. In later studies, however, disc formation was never again observed in pure cultures of this

CHAPTER 3.2.11

morphotype, so that it was concluded that the presence of this trait within the genus Leptothrix was confined to the species L. lopholea (Mulder, 1989a). If the authors of the abovementioned studies have in fact described the same species, the ability to form discs would not distinguish between L. discophora and L. lopholea, this trait being more stable in the latter. To avoid this kind of confusion, descriptions of new strains should be based on freshly isolated pure cultures that were studied, if possible, under conditions most similar to natural habitat conditions. Lack of Reference Strains A major problem in the taxonomy of the genus Leptothrix is the availability of reference strains. The type species of the genus Leptothrix, L. ochracea, is not cultured and the description is based only on morphological observations. Although the description of “L. pseudo-ochracea” was based on pure cultures, no type strain was designated. On the other hand, the type strains of L. lopholea, LVMW 124, and L. cholodnii, LVMW 99, are not available from public culture collections and have apparently been lost. The strain LMG 7171 can be used as reference strain for L. cholodnii until a neotype is designated, but for L. lopholea, no other strains are available, preventing detailed taxonomic studies on this species.

Ecology Ecology of Sheath Formation The production of a sheath takes place at the expense of energy and requires synthesis and excretion of a large amount of cellular material. Therefore, it can be assumed that sheath formation was developed by microorganisms under a selective pressure and offers several ecological and nutritional advantages in the environment. A linear arrangement of single cells within a tubular sheath enables bacteria to form filaments, without actual enlargement of cell size. It has been shown that filamentous growth is an effective strategy of bacteria to exceed the size limit of particles edible by protozoa, thereby allowing them to escape from grazing (Sommaruga and Psenner, 1995). In addition, sheaths provide cells with physical protection from infection by bacteriophages (Winston and Thompson, 1979), bacterial predators (Venosa, 1975), and bacterivorous metazoa. Obviously tough sheaths impregnated with ferric oxides cannot be consumed by metazoa and are not penetrated by bdellovibrios or phages. Rigid sheaths in combination with extracellular slime represent an ideal matrix for the rapid

CHAPTER 3.2.11

The Genera Leptothrix and Sphaerotilus

build-up of biofilms in the form of floating aggregates or dense layers on solid surfaces. The ecological advantages of biofilm formation for bacteria are well known and include reduction of flow rates in running waters, adsorption of nutrients, and establishment of heterogenous microniches characterized by various chemical gradients. The reversible inhibition of sheath formation by high nutrient concentrations in some Sphaerotilus strains may be an indication for the important role played by the sheath surface in the accumulation of nutrients (Gaudy and Wolfe, 1961). The sheath provides cells with an additional compartment close to their outer surface. Enzymes and proteins which are secreted by the cell can be incorporated in the sheath matrix, thereby allowing the extracellular production or degradation of compounds. It is possible that members of both genera use the sheath or the space between sheath and outer cell surface for the oxidation of iron and/or manganese to avoid toxic concentrations of metal compounds within the cell.

Methods for Studying Distribution and Abundance Members of the Sphaerotilus-Leptothrix group are not only abundant in natural environments, but also may play a role in industrial processes leading to several technical problems. They were associated with bulking of activated sludge (Wagner et al., 1994), corrosion of stainless steel (Olesen et al., 2000), clogging of water distribution systems (Dondero, 1975), and slime formation in paper mill factories (Pellegrin et al., 1999). To prevent damage caused by the prolific growth of these organisms, it could be advantageous to

763

monitor them in environmental samples without time-consuming enrichment and cultivation steps. Although in some samples the affiliation of bacteria to the Sphaerotilus-Leptothrix group may be assigned on the basis of phase-contrast microscopic appearance, distinction between genera or single species is hardly possible. Consequently, culture-independent methods for the identification and detection of these species were developed. Two approaches could be promising: whole cell hybridization with fluorescently labeled oligonucleotide probes and polymerase chain reaction (PCR) detection of Leptothrix species using primers targeting mofA, an enzyme gene involved in manganese oxidation. Whole Cell Hybridization Fluorescence in situ hybridization (FISH) using oligonucleotide probes is a well established technique for the identification and in situ detection of microorganisms based on signature regions of their 16S rRNA genes (Amann et al., 1995). Several probes were developed targeting 16S rRNA sequences of members of the SphaerotilusLeptothrix group. A check of published probe sequences against a current database of 16S rRNA gene sequences (Ludwig and Strunk, 1997) revealed, however, that some probes which were originally intended to be specific for a single species are in fact targeting a variety of different species, partly not even belonging to the Sphaerotilus-Leptothrix group. To minimize the risk of mistaken identification of uncultured bacteria by crossreactivity of probes, it is advisable to use simultaneously at least two different specific probes, distinguishable by fluorescence label. Suitable combinations of probes for the Sphaerotilus-Leptothrix group can be found in Table 3.

Table 3. Specificity of oligonucleotide probes targeting 16S rRNA sequences of members of the Sphaerotilus-Leptothrix group. Hybridization with probea

Sequence of: Species Leptothrix cholodnii

Strain

Accession no.

CCM 1827 X97070 SP-6 L33974 L. discophora SS-1T L33975 L. mobilis Feox-1T X97071 Sphaerotilus sp. IF4 AF072914 S. natans 6T L33980 Target region of probe (E. coli positions) Sequences with complementary target region, not affiliated to the Sphaerotilus-Leptothrix groupd

SNAb

PSP-6c

LDIb

PS-1c

+ + − + + + 656–673 13

+ + − + − − 138–155 9

− − + − − − 649–666 4

+ + + − − − 66–82 0

Symbols: +, hybridization of probe; −, no hybridization of probe under stringent conditions; and T, type strain. a Reactivity of the probe was either checked experimentally or determined by sequence comparison. b Data from Wagner et al. (1994). c Data from Siering and Ghiorse (1997). d Only sequences of cultured strains available in public data bases were counted.

764

S. Spring

CHAPTER 3.2.11

The identification of distinct species by FISH is still difficult, even by using a combination of specific probes, but the combined observation of morphological features (e.g., sheath formation) and hybridization signals should at least enable the identification of uncultured bacteria at the genus level. In Figure 2, the in situ detection of uncultured Leptothrix bacteria in an environmental wetland sample is shown. Low signal intensities of labeled cells and samples with low numbers of target cells represent frequently encountered problems of the FISH method. In habitats with low numbers of sheathed bacteria, e.g., water distribution systems or sediments, the application of microscope slides can be a promising approach to obtain suitable samples for hybridization experiments. Microscope slides were successfully used for the enrichment of sheathed bacteria by employing to advantage their ability to adhere efficiently to smooth surfaces (Spring et al., 1996; Carlile and Dudeney, 2001). After removal and cleaning, the overgrown side of these slides can be used for FISH experiments. To improve signal intensities of labeled cells, it may help to treat samples prior to hybridization with diluted hydrochloric or oxalic acid, to remove iron and manganese oxides from sheaths, which could otherwise pre-

a

vent efficient penetration of oligonucleotide probes into cells. Diagnostic PCR An alternative approach for the detection of Leptothrix bacteria was developed by Siering and Ghiorse (1997b). They used specific PCR primers for the amplification of the gene mofA, encoding a putative multi-copper oxidase involved in manganese oxidation by Leptothrix species (see section “Oxidation of Fe2+ and Mn2+” in this Chapter). The gene mofA was originally retrieved by cloning a DNA fragment of Leptothrix discophora SS-1, but homologous genes were identified also in several other Leptothrix species (Siering and Ghiorse, 1997b). The application of nondegenerate PCR primers enabled the in vitro amplification of a 706-bp portion of the gene mofA from a pure culture of L. discophora SS-1 and extracted nucleic acids from a water sample of a wetland iron seep, but not from nucleic acids extracted from a sediment sample of the same site. The obtained results were in agreement with previous observations which indicated that in this habitat, the growth of Leptothrix bacteria is restricted to ferromanganese surface films and the root zone of Lemna species in the water column. Potential advantages of this protocol are an increased specificity and sensitivity compared to the FISH method. However, in contrast to the latter method, no information on the morphology of the detected cells can be obtained.

The Genus Leptothrix Habitats

10 mm b

Fig. 2. Laser scanned epifluorescence (a) and matching laser scanned phase contrast image (b) of previously unidentified filamentous bacteria labeled by the PS-1 oligonucleotide probe in a sample of particulate material from the Sapsucker Woods wetland, Ithaca, NY. From Ghiorse et al. (1996), with permission.

Leptothrix species are widely distributed in the environment and can be easily found at sites which are characterized by a circumneutral pH, an oxygen gradient and a source of reduced iron and manganese minerals. Typical habitats include iron seeps of freshwater wetland areas, forest ponds, iron springs and the upper layers of sediments (Ghiorse and Ehrlich, 1992). At suitable sites, the massive growth of these organisms can be easily observed with the naked eye as ocherous masses emerging as surface films, solid mats or fluffy, dispersed material. Depending on the amount of oxidized manganese, the color can vary from yellowish-orange to dark brown. A wetland iron seep, shown in Fig. 3, is characterized by a pronounced ocherous color originating from a prolific growth of Leptothrix bacteria. An iron-oxidizing microbial mat that developed at an iron seep where anoxic groundwater (rich in ferrous iron) flows over a stone wall was described in detail by Emerson and Revsbech (1994). They could demonstrate that Leptothrix

CHAPTER 3.2.11

The Genera Leptothrix and Sphaerotilus

7

6

765

5 4

8 3 9

Fig. 3. Ellis Hollow iron seep (near Ithaca, NY). A wetland site with massive growth of Leptothrix species (mainly L. ochracea) resulting in typically colored surface films. From Ghiorse and Ehrlich (1992), with permission.

species represent a significant fraction of the active biomass in the upper few millimeters of this dense microbial mat. Further studies presented indirect evidence for the association of Leptothrix bacteria with lacustrine ferromanganous micronodules (Stein et al., 2001) or microbial mats in areas of possible deep-lake hydrothermal venting (Dymond et al., 1989). Most Leptothrix species are restricted to natural, unpolluted environments with low concentrations of easily degradable organic nutrients. A prominent exception is L. cholodnii, which is frequently detected in sewage sludge (Kämpfer, 1997).

Isolation Several methods for the enrichment of Leptothrix species from the environment are based on the tendency of these filamentous organisms to attach to surfaces. Mulder and Van Veen (1963) used continuous flow devices to imitate the natural growth conditions of these organisms. A schematic of such an apparatus is shown in Fig. 4. For producing iron-containing ditch water, an iron-cylinder is filled with iron-stone soil supplemented with 1 to 2 g of ferric carbonate/kg of soil. After an incubation period of about 3 weeks, during which the soil is kept saturated with water, enough ferric iron is reduced to ferrous iron to start a continuous flow of tap water percolating through the soil. The soil extract running off the iron cylinder from the upper outlet is sterilized by filtration and supplied dropwise to the upper Erlenmeyer flasks, which can be inoculated with various samples containing Leptothrix bacteria. To avoid rapid nonbiological oxidation of ferrous iron, the upper Erlenmeyer flasks can be aerated with a gas mixture of low oxygen content. Comparable to the situation in

2

1

2

10 Fig. 4. Apparatus for growing Leptothrix species in running artificial iron-containing ditch water. (1) Inlet for tap water. (2) Distributor for the incoming tap water. (3) Cylinder containing iron-stone soil. (4) Seitz filter. (5) Manifold for distributing the sterile iron-containing water. (6) Inlet for the gas mixture (1% O2, 5% CO2, and 94% N2). (7) Inoculation tube. (8) Erlenmeyer flask at a high level. (9) Erlenmeyer flask at a low level fitted with outlets (10). Reproduced with permission from Mulder and Deinema (1992).

unpolluted slowly running water, bacteria which are able to attach to solid surfaces have a selective advantage within this device because they obtain more nutrients than bacteria moving with the medium. The described apparatus can be also applied for the observation of pure cultures of Leptothrix bacteria under conditions similar to their natural environment. A less laborious enrichment method was introduced by Rouf and Stokes (1964), who filled glass cylinders with water taken from the environment and added extracted alfalfa straw as nutrient source, MnCO3, and freshly precipitated Fe(OH)3. After several days, the flocculent growth of sheathed bacteria adhering to the walls of the cylinder indicated the enrichment of Leptothrix bacteria. An alternative method, which is also based on the capability of Leptothrix species to attach to smooth surfaces, was applied by Spring et al. (1996). Microscope slides were put into a sample of freshwater sediment, and after several weeks, sheaths of filamentous bacteria encrusted with ferric oxides covered parts of the slides. The slides can be removed from the sediment and used for isolation after washing in sterile tap water to remove loosely attached sediment bacteria. Isolation of pure cultures can be achieved by streaking material from enrichment cultures on previously dried agar plates containing low levels of nitrogen and carbon sources. Enrichment cul-

766

S. Spring

CHAPTER 3.2.11

tures may not be necessary if natural environments are studied in which Leptothrix bacteria can be detected by their flocculent growth. Flocculent cell material from these sites can be washed several times with sterile tap water and streaked directly on solid media. The isolation medium used by Rouf and Stokes (1964) has the following composition (per liter of tap water): Peptone, 5.00 g; ferric ammonium citrate, 0.15 g; MgSO4 ·7 H 2O, 0.20 g; CaCl2, 0.05 g; MnSO4 ·H 2O, 0.05 g; FeCl3 ·6 H2O, 0.01 g; and agar, 12.00 g. The plates are incubated at 25°C for 5–14 days in the dark. Colonies of Leptothrix strains can be easily distinguished from most contaminating bacteria on this medium by their dark-brown color.

Identification So far six different species of Leptothrix can be distinguished on the basis of phenotypic characteristics. Reference strains for taxonomic studies are only available for three species, viz., L. cholodnii, L. discophora and L. mobilis. Descriptions of the remaining species are incomplete because they are based solely on microscopic observations (L. ochracea) or poorly characterized isolates (L. lopholea and “L. pseudoochracea”), hence preventing an accurate identification of newly isolated strains using culture-dependent methods. The name of “Leptothrix pseudo-ochracea” was not included in the Approved Lists of Bacterial Names (Skerman et al., 1980) and has therefore no formal taxonomic status. Differential characteristics of the known

Leptothrix species are listed in Table 4. Detailed morphological descriptions follow below. L. OCHRACEA Leptothrix ochracea is the most common iron-precipitating ensheathed bacterium that probably occurs all over the world in slowly running ferrous iron-containing waters, poor in readily decomposable organic matter. Low oxygen tensions seem to be required for prolific growth of this species (Emerson and Revsbech, 1994). The pronounced development and activity of L. ochracea in iron- and manganese-containing waters give rise to the accumulation and deposition of large masses of ferric oxide and, probably, manganese dioxide (MnO2), which are thought to be responsible for the formation of bog ore (see for instance, Ghiorse and Chapnick, 1983). Descriptions of this species are based on observations in the natural environment or in the laboratory on enrichment cultures of slowly running soil extract. The most typical characteristic of L. ochracea is the formation of large numbers of almost empty sheaths within a relatively short time. The mechanism of this procedure can be followed in a slide culture of the organism, in an iron-containing soil extract medium, under a phase-contrast microscope. In this way, the behavior of L. ochracea in crude culture can be observed continuously. It will be seen that chains of cells leave their sheath at the rate of 1–2 µm/ min, continuously producing a new smooth and hyaline sheath connected with the old envelope (Fig. 5a). Impregnation and covering of the sheaths with iron probably take place after the

Table 4. Differential characteristics of Leptothrix species. Characteristic Cell dimensions Width (µm) Length (µm) Flagella Monotrichous polar Polytrichous subpolar Holdfasts False branching Deposition of MnO2 on sheath surface Growth at 35°C pH 8.5 Growth ona d-Fucose Fumarate dl-Lactate G+C content (mol%)

L. ochracea

“L. pseudo-ochracea”

L. lopholea

L. cholodnii

L. discophora

L. mobilis

1.0 2–4

0.8–1.3 5–12

1.0–1.4 3–7

0.7–1.5 2.5–15

0.6–0.8 2.5–12

0.6–0.8 1.5–12

+ − − − −

+ − − − +

− + + + +

+ − − − +

+ − V + +

+ − − − ND

ND ND

ND ND

ND ND

+ +

− −

+ +

ND ND ND ND

ND ND ND ND

ND ND ND ND

− − + 68–70

+ − − 71

− + − 68

Symbols: +, present in all strains; −, absent in all strains; ND, not determined; and V, variable (depending on growth conditions). a Growth was determined in GMBN medium supplemented with the respective carbon source (Kämpfer et al., 1995).

CHAPTER 3.2.11 Fig. 5. Morphological characteristics of L. ochracea. Bar = 10 µm. (a) Phase contrast micrograph of cells moving out of sheaths and subsequently forming new sheaths. From Van Veen et al. (1978), with permission. (b) Broken old sheaths covered and impregnated with ferric hydroxide in slowly running iron(II)-containing soil extract. From Mulder and Van Veen, 1963, with permission. (c, d) Acridine orange staining of cells partly covered by a hyaline sheath. Same field viewed by differential interference contrast (c) and epifluorescence microscopy (d). Intact bacterial cells stained with acridine orange fluoresce bright green under violet illumination. Note that the chain of single cells appears as continuous filament when viewed by interference contrast. Courtesy of W. C. Ghiorse.

The Genera Leptothrix and Sphaerotilus a

c

cells have left the envelopes. Aged golden-brown and highly refractile sheaths are brittle, so that they are easily broken into relatively short fragments. Viewed under phase contrast, such fragments are very characteristic and appear similar to broken glass capillaries (Fig. 5b, c). In Mn(II)containing environments or enrichment cultures, sheaths of this species can be easily distinguished from sheaths of the other Leptothrix species which are characterized by irregular encrustations of granular MnO2. In contrast, the sheaths of L. ochracea are only impregnated with ferric oxides and lack MnO2 deposits, leading to a smooth and slender appearance of aged sheaths. The lack of MnO2 encrustations in sheaths of L. ochracea can be detected either by using an electron microscope equipped with an X-ray energy dispersive microanalysis system (Ghiorse and Ehrlich, 1992) or by treatment with oxalic acid (Carlile and Dudeney, 2001). A diluted solution (1%) of oxalic acid, a solvent for hydrated ferric oxide, can make the sheaths of L. ochracea transparent and almost disappear, whereas sheaths of other Leptothrix species are unaffected owing to the impregnation with manganese oxides. Investigators who have studied and described L. ochracea under natural conditions were unable to obtain pure cultures (Cholodny, 1926; Charlet and Schwartz, 1954). Others who thought they had isolated L. ochracea had, in fact, described one of the other species of this genus (Winogradsky, 1888; Winogradsky, 1922; Molisch, 1910; Lieske, 1919; Cataldi, 1939; Präve,

767

b

d

1957). In some instances, an organism resembling L. ochracea has been isolated, viz., “L. pseudo-ochracea.” A potential lithoautotrophic metabolism of L. ochracea with Fe(II) as electron donor could, therefore, never be proved with pure cultures and is uncertain, since the organism normally grows at a pH value of 6–7, at which Fe(II) is readily oxidized nonbiologically. In addition, its Mn (II)-oxidizing capacity, which probably occurs under natural conditions, has never been confirmed. It is, however, possible that in this species, the Mn(II) oxidizing factors are secreted into the surrounding environment leading to the precipitation of MnO2 granules away from the sheath surface. “L. PSEUDO-OCHRACEA” Cells are more slender than those of the other Leptothrix species (Table 4), and are very motile by one thin polar flagellum. Even chains of 6–10 cells may show an undulatory locomotion after leaving their sheath. This characteristic may account for the relatively large number of empty sheaths in culture, compared with the number found in cultures of most other Leptothrix species; however, L. ochracea possesses even more empty sheaths. In slowly flowing ferrous iron-containing soil extract, the sheaths become impregnated with ferric oxide and appear yellow-brown. In this respect, the organism resembles L. ochracea. However, in media with added manganese compounds, the sheaths are covered with small granules of MnO2, enabling an easy distinction from

768

S. Spring

a

the sheaths of L. ochracea which are not impregnated with manganese oxides. On Mn(II)-containing agar, the black-brown colonies are very filamentous and may exceed a width of 10 mm. On basal agar media containing 0.1% peptone and 0.1% glucose, the organism may grow in concentric rings. The normal habitat of “L. pseudo-ochracea” is the slowly running, unpolluted, iron- and manganese-containing freshwater of ditches and brooklets. This species may also be found in slightly polluted water. L. LOPHOLEA Leptothrix lopholea resembles S. natans to a greater extent than do the other Leptothrix species. It produces polytrichous subpolar flagellation and forms holdfasts and false branches. Strains may grow also in rich media. Cells usually develop short-sheathed filaments radiating from a cluster of holdfasts, giving rise to many tiny flocs when the cells are grown in liquid media (Fig. 6). Deposition of iron and manganese oxides is more pronounced on holdfasts than on filaments. On Mn(II)-containing agar media, encrustation of sheaths with MnO2 is retarded, so that colonies at first are white and later become blackbrown. Cell growth responds poorly to an increased supply of organic nutrients. Strains that do show a good response oxidize manganese more slowly. This species may be isolated from slowly flowing, unpolluted or polluted freshwater and from activated sludge. L. CHOLODNII Cells of freshly isolated strains are usually found in long chains inside the sheaths. Single motile cells may be seen outside the sheaths. In the presence of Mn(II), the

CHAPTER 3.2.11 b

Fig. 6. Leptothrix lopholea. (a) Bacterial flocs with black-brown MnO2 deposits. Reproduced with permission from Mulder and Deinema (1992). (b) Many trichomes radiating from common holdfasts. Bar = 10 µm. From Van Veen et al. (1978), with permission.

sheaths become covered with granular MnO2 (Fig. 7a). At some sheath locations, the MnO2 deposits may even exceed 10 µm. Leptothrix cholodnii, in contrast to other Leptothrix species, responds to an increased supply of organic nutrients (Table 2). This results in relatively large colonies (up to 5 mm in diameter) on nutrient-rich agar media. On Mn(II)-containing agar, blackbrown hairy colonies are formed (Fig. 7b), particularly when the organism is seeded densely. Most strains display a strong tendency to dissociate spontaneously and to produce smooth rather than the typical rough colonies (Fig. 7c). Such mutant strains are largely sheathless and oxidize manganese slightly or not at all (Mulder and Van Veen, 1963; Rouf and Stokes, 1964; Stokes and Powers, 1965). In agreement with its nutritional requirements, L. cholodnii is found in slowly running iron- and manganese-containing unpolluted waters or in polluted waters, particularly in activated sludge. L. DISCOPHORA Cells are relatively small compared to those of the other Leptothrix species described (Table 4). They may occur in narrow sheaths or be free-swimming; free cells are motile by a thin polar flagellum at one or both poles. The manganese-oxidizing and ferric oxidestoring capacities of this organism are very pronounced. In the presence of Mn(II), the sheaths are heavily but irregularly encrusted with MnO2, giving rise to sheaths of sometimes 10 µm thickness. Under natural conditions, holdfasts may be formed and sheaths are covered with a slime capsule which tapers toward the growing tip and is impregnated with hydrated ferric oxides (Carlile and Dudeney, 2001). In enrichment media with both manganese (II) and iron (II), as in slowly

CHAPTER 3.2.11 Fig. 7. Leptothrix cholodnii. (a) Sheaths encrusted with MnO2. Bar = 10 µm. Reproduced with permission from Mulder and Deinema (1992). (b, c) Colony morphologies. (b) Filamentous colony on agar with MnCO3 (medium 1, Mulder and Van Veen, 1963). Bar = 100 µm. Reproduced with permission from Mulder and Deinema (1992). (c) Smooth colony on the agar medium of Rouf and Stokes (1964). Bar = 500 µm.

The Genera Leptothrix and Sphaerotilus a

769 b

c

flowing soil extract, the sheaths become covered with a thick, dark brown, fluffy layer of ferric oxide and MnO2 which may increase the diameter of the trichomes up to about 20–25 µm (Fig. 8a). Following isolation, the ability to form sheaths is easily lost in this species (Adams and Ghiorse, 1986). False branching is regularly observed, even with sheathless strains (Fig. 8b). In older cultures, coccoid bodies and cell evaginations are formed. Colonies on the solid medium of Rouf and Stokes (1964) are about 1 mm in diameter, more or less circular in shape, flat, and darkbrown (Fig. 8c). Under certain conditions, filamentous colonies may be formed. Increasing the supply of nutrients such as glucose, peptone, methionine, purine bases, vitamin B12, biotin and thiamine only increases growth slightly. Visible aggregates are formed when grown in liquid media. The normal habitat is slowly running, unpolluted, iron- and manganese-containing water of ditches, rivers or ponds. L. MOBILIS Cells are similar in width to L. discophora cells (Fig. 9a), but are usually shorter in

length. False branching, which is typical for L. discophora, is absent in cultures of L. mobilis. Cells are highly motile by a single polar flagellum. Sheaths are not formed under laboratory conditions. Colonies on the agar medium of Rouf and Stokes (1964) are about 1 mm in diameter, circular sometimes with frayed edges, flat, smooth and dark-brown (Fig. 9b). Visible aggregates are formed when grown in liquid media. The species description is based on only one strain isolated from the sediment of a freshwater lake (Spring et al., 1996).

Preservation Stock cultures can be stored on agar slants of the medium of Rouf and Stokes (1964) for about two months at 4°C. Rouf and Stokes (1964) reported a better survival of their strains if stored at room temperature instead of refrigerated. Most Leptothrix strains do not survive lyophilization. For the long-term preservation of these strains, freezing in liquid nitrogen is recommended using suspensions of cells in freshly prepared medium supplemented with 5% dimethyl sulfoxide (DMSO) as cryoprotectant.

770

S. Spring

CHAPTER 3.2.11

a

b

Fig. 8. Leptothrix discophora. (a) Sheaths covered with ferric hydroxide and manganese dioxide in running iron(II)- and manganese(II)containing soil extract. Bar = 10 µm. From Mulder and Van Veen (1963), with permission. (b) Cells of a sheathless strain showing false branching. Bar = 10 µm. From Spring et al. (1996), with permission. (c) Smooth colony on MnSO4-containing agar. MnO2 is present within the colony and in a halo containing no bacteria. Bar = 100 µm. From Mulder and Van Veen (1963), with permission.

b

Fig. 9. Leptothrix mobilis. (a) Dimensions of single cells grown in the medium of Rouf and Stokes (1964). Bar = 10 µm. From Spring et al. (1996), with permission. (b) Colony morphology on agar medium of the same composition. Dark-brown, granular deposits of MnO2 are visible in the frayed edge of the colony. Bar = 500 µm.

c

a

Physiology The genus Leptothrix is characterized by two remarkable metabolic activities: Formation of sheaths and oxidation of iron and manganese. Oxidation of Fe2+ and Mn2+ It could be demonstrated in several studies that the deposition of metal-oxides in Leptothrix species is biologically controlled. Oxidation of manganese and iron is catalyzed by various metal-oxidizing factors actively secreted by the cell (De Vrind-de Jong et al., 1990). At circumneutral pH, biological oxidation of iron is difficult to distinguish from the chemical oxidation by oxygen. Only with the identification of an iron-oxidizing protein with a molecular

weight of 150 kDa in spent culture medium of the sheathless strain Leptothrix discophora SS-1, the capability of biological iron-oxidation in Leptothrix species could be clearly demonstrated (Corstjens et al., 1992). The function of ironoxidation is still unknown, but it seems unlikely to play a role in the generation of energy. It has to be noted, however, that Gallionella ferruginea, which is also neutrophilic and microaerophilic, thrives in the same habitats as L. ochracea and is able to gain energy from chemolithoautotrophic iron-oxidation (Hallbeck et al., 1993). Several manganese-oxidizing factors could be identified in culture supernatants of Leptothrix species. One component with an estimated molecular weight of 110 kDa could be purified and partly characterized. It is called “manga-

CHAPTER 3.2.11

nese-oxidizing factor” (MOF) and consists of protein and probably polysaccharide (Emerson and Ghiorse, 1992). It is assumed that this component is part of a larger complex that originates from membranous blebs and is associated with the sheath structure (Brouwers et al., 2000). Antibodies raised against the MOF protein of L. discophora SS-1 allowed retrieval of the gene mofA, encoding a putative multi-copper oxidase. A strong indication for an active role of this copper-dependent enzyme in manganeseoxidation is the observation that addition of small amounts of copper to actively growing cultures enhances the rates of manganese oxidation considerably (Brouwers et al., 2000). In addition to multi-copper oxidases, c-type hemes could be involved in metal-oxidation by Leptothrix species. Genes encoding proteins with potential heme-binding sites were identified as part of the operon encoding mofA. So far, the beneficial effects of manganese oxidation are largely unknown, but it is unlikely that the deposition of manganese oxides has been developed without conferring an important advantage to these bacteria. The benefits of manganese oxidation are probably so difficult to determine because they are only effective under natural conditions difficult to imitate in the laboratory. Traditionally, it was assumed that the positive effects of Mn(II)-oxidizing enzymes and manganese oxides are based on the detoxification of harmful oxygen species (superoxide and peroxide) or on the adsorption of toxic compounds (e.g., heavy metals). Recently, an interesting new perspective was introduced by Sunda and Kieber (1994). They found that manganese oxides are strong chemical oxidants able to attack complex organic compounds (e.g., humic substances), thereby leading to the release of small organic molecules which can be easily assimilated by bacteria. Structure and Composition of the Sheath The overall structure and chemical composition of sheaths formed by Leptothrix species resemble those of Sphaerotilus natans, although several distinguishing traits were found. One characteristic which can be used for the differentiation between both genera is the structure of the sheath surface. In electron micrographs of unstained preparations, the sheath surface appears rough in Leptothrix species and smooth in strains of Sphaerotilus (Figs. 10 and 12b). Emerson and Ghiorse (Emerson and Ghiorse, 1993a; Emerson and Ghiorse, 1993b) performed a detailed investigation of the ultrastructure and chemical composition of the Leptothrix sheath with the sheath-forming strain L. cholodnii (formerly “L. discophora”) SP-6. They reported that it consists of a condensed fabric of 6.5-nm-

The Genera Leptothrix and Sphaerotilus

771

Fig. 10. Electron micrograph of a Leptothrix sheath showing a rough surface. Bar = 1 µm. Reproduced with permission from Mulder and Deinema (1992).

diameter fibrils underlying a more diffuse outer capsular layer. The inner sheath layer has a thickness of 30–100 nm and seems to be associated with the outer layer of the Gram-negative cell wall by membrane evaginations. The purified sheath substance contains approximately 34% polysaccharide, 24% protein, 8% lipid, and 4% inorganic material. As major components of the polysaccharide moiety, uronic acids and amino sugars, which are probably in the N-acetylated form, were identified, whereas neutral sugars could not be detected. The sheath proteins are rich in cysteine residues (6 mol%), which confer numerous disulfide- and sulfhydryl-groups to the sheath. The heteropolysaccharide and protein moieties appear to be tightly associated or connected. On the basis of these findings, it was concluded that the principal structural elements of the sheath are proteoglycan fibrils that are covalently linked to each other by interfibril disulfide bonds resulting in a stable fabric. It is possible that the difference between the condensed inner layer and diffuse capsular layer is caused by an increased amount of free sulfhydryl groups in the exterior sheath-layer. A further characteristic of the sheath is its negative charge due to free carboxyl groups, originating mainly from the uronic acids of the sheath-proteoglycans. Both chemical groups, the free sulfhydryl and the free carboxyl groups, provide the sheath with numerous sites for binding of metal cations, especially Mn2+ and Fe2+. According to the working model

772

S. Spring

CHAPTER 3.2.11 Mn oxidizing protein(s) Exterior layer Interior layer

bleb PHA

Polyphosphate

COOS

SH

Cells

SH S S

S

Fig. 11. Schematic model of the architecture and potential function of the sheath in L. cholodnii SP-6. PHA, polyhydroxyalkanoate. From Emerson and Ghiorse (1993a), with permission.

S S

COO-

COO-

presented in Fig. 11, the anionic sheath-fibrils capture and concentrate soluble Mn(II) ions within the sheath, where they can be efficiently oxidized by excreted manganese-oxidizing factors associated with the sheath (Emerson and Ghiorse, 1993a). Manganese- and iron-oxidizing factors are probably excreted from the cell and transported to the sheath as membranous blebs containing protein complexes consisting of multi-copper oxidases and cytochromes (Emerson and Ghiorse, 1992).

The Genus Sphaerotilus Habitat Members of the genus Sphaerotilus are part of the natural microbial community in slowly running freshwater streams, ditches and ponds (Stokes, 1954). In contrast to Leptothrix sp., massive growth is typical in freshwater streams that receive high levels of organic pollution from sewage or industrial wastes, especially from paper, potato, dairy or other agricultural industries. When Sphaerotilus becomes established in a polluted river, it frequently proliferates so extensively that it lines the riverbed for long distances with a wooly, filamentous carpet (Dondero, 1975). In poor settling activated sludge (so-called “bulking sludge”), Sphaerotilus is frequently found among other filamentous microorganisms (Eikelboom, 1975). Bulking or foaming of activated sludge is a common problem in water purification plants and prevents a ready separation of bacterial sludge flocs from the treated water. It appears that conditions which stimulate the growth of Sphaerotilus natans and/or other fila-

mentous bacteria favor also the bulking of activated sludge. The correlation of conditions which favor the prolific growth of a distinct bacterial species with sludge bulking is however hardly possible owing to the high phenotypic diversity of filamentous bacteria which may be involved in this phenomenon (Kämpfer, 1997). The uncontrolled growth of Sphaerotilus in artificial environments bears several risks of economic importance. It can contribute pyrogenic material to purified water for medical injection (Dondero, 1975) and cause damage to technical equipment or machines (Pellegrin et al., 1999). On the other hand, beneficial effects of the growth of S. natans are also discussed, e.g., the biosorption of heavy metals in wastewater at low pH values (Solisio et al., 2000; Esposito et al., 2001).

Isolation Enrichment Procedures In many environments sheathed bacteria occur only in low numbers, for instance in activated sludge or nonpolluted water samples. The use of of enrichment cultures may facilitate the successful isolation of Sphaerotilus sp. from such habitats. For that purpose, variations of Winogradsky’s hay infusion technique have been used (Mulder and Deinema, 1992). Extracted alfalfa straw (Stokes, 1954) or extracted pea straw (Mulder and Van Veen, 1963) serve as the nutrient material. Most of the soluble organic matter should be removed to prevent proliferation of undesirable organisms. This can be achieved by boiling and extracting the straw after it has been cut into small pieces. Suspensions of the extracted straw in tap water (1–8% [w/v]) are distributed in Erlenm-

CHAPTER 3.2.11

eyer flasks and used as enrichment medium. After inoculation with 5–20% of a water sample or activated sludge and incubation for about one week at 22–25°C, filaments of Sphaerotilus may be seen in the medium upon microscopic observation. Often tufts of filaments can be found attached to the pieces of straw and also to the side of the flasks at or near the surface of the medium, thereby enabling easy removal using glass capillaries. Pure cultures can be obtained from homogenized portions of the enriched material as described in the following paragraph. Direct Isolation Often slimy masses of Sphaerotilus are found attached to submerged surfaces in polluted, slowly running water. Material from such sites can be successfully used for the direct isolation of S. natans or related strains by streaking on agar plates. To remove contaminating cells, the collected material should be washed with sterile tap water several times. Homogenization of the washed flocs by blending for a very short time may be useful. However, activated sludge flocs containing many filaments of Sphaerotilus should be streaked directly on the agar plates without homogenization step to avoid release of numerous, nonfilamentous cells by destroying the floc structure. The agar plates used should be dry and contain only low levels of nitrogen and carbon to limit the size of undesirable bacterial colonies, leaving large areas for the filamentous organisms. A further reason for the use of nutritionally poor media is the observation that Sphaerotilus often forms smooth colonies on nutrient-rich agar, instead of the typical rough colonies, which can be easily recognized. To inhibit the growth of contaminating fungi, the agar medium can be supplemented with cycloheximide (0.005 g ·liter –1; Pellegrin et al., 1999). A suitable isolation medium was proposed by Mulder (1989b), having the following basal composition per liter of distilled water: KH2PO4, 27 mg; K2HPO4, 40 mg; Na2HPO4 ·2 H 2O, 40 mg; CaCl2, 50 mg; MgSO4 ·7 H 2O, 75 mg; FeCl3 ·6 H2O, 10 mg; MnSO4 ·H 2O, 5 mg; ZnSO4 ·7 H 2O, 0.1 mg; CuSO4 ·5 H 2O, 0.1 mg; Na2MoO4 ·2 H2O, 0.05 mg; cyanocobalmin, 0.005 mg; peptone, 1 g; glucose, 1 g; and agar, 7.5 g. Upon inoculation and incubation of these plates at 20–30°C, colonies of Sphaerotilus may be seen and tentatively identified within a few days by their characteristically flat, dull, cottonlike appearance. Confirmation of the identification may be achieved by microscopic observation. Sphaerotilus may also be isolated by spread plate techniques with or without a previous centrifugation or homogenization step on a variety

The Genera Leptothrix and Sphaerotilus

773

of agar media (Eikelboom, 1975; Williams and Unz, 1985; Ziegler et al., 1990), including the commercially available R2A agar (Seviour et al., 1994).

Identification The genera Sphaerotilus and Leptothrix are closely related and share many phenotypic characteristics. Nevertheless, a clear differentiation of both genera is possible on the basis of phenotypic traits summarized in Table 2. The only recognized species of Sphaerotilus is currently S. natans. Typical morphological characteristics of this species are false branching of filaments, extensive slime production, attachment of filaments to solid surfaces by holdfast formation (Fig. 12a), deposition of hydrated ferric oxides on sheaths, and motility of cells by means of a bundle of subpolar flagella (Fig. 12b). In contrast to those of Leptothrix, the sheaths of Sphaerotilus occurring in natural habitats are usually thin and hyaline without encrustations by ferric or manganese oxides (Fig. 12c). However, the capability of Fe(II) oxidation can be easily demonstrated in this species by cultivation in media containing soluble iron compounds and low nutrient concentrations, e.g., soil extract enriched with ferrous iron. Under these growth conditions, the sheaths of S. natans turn yellowbrown and resemble in appearance those of Leptothrix ochracea. Most of the pronounced morphological characteristics of S. natans are largely dependent on the strain and cultivation conditions. Sheath formation can be inhibited by high levels of nutrients in the medium—peptones being more effective than carbohydrates— resulting in the formation of smooth instead of rough, filamentous colonies (Mulder and Van Veen, 1963). This loss of sheath-forming capability is reversible and can occur also spontaneously. Growth of pure cultures in liquid media is usually flocculent, but sometimes pellicular or homogenous (Pellegrin et al., 1999). Upon prolonged incubation, large, circular bodies resembling protoplasts may appear in broth cultures. Their formation is probably due to the production of enzymes involved in the decomposition of cell walls during the death phase (Phaup, 1968). The nutritional versatility of S. natans and related strains is remarkable. They can utilize a variety of carbon and nitrogen sources, tolerate a wide range of nutrient concentrations and can grow under low partial pressures of oxygen. Utilization of fructose, glucose, maltose, sucrose, lactate, pyruvate, and succinate as sole sources of carbon was reported by Stokes (1954), Mulder and Van Veen (1963), Kämpfer (1998), and Pellegrin et al. (1999). Numerous other carbon

774

S. Spring

CHAPTER 3.2.11 a

b

c

Fig. 12. Sphaerotilus natans. (a) Cells with internal granules of poly-β-hydroxybutyrate (white arrowhead) and filaments showing holdfasts (black arrowheads). Bar = 10 µm. (b) Electron micrograph of a single cell with a subpolar tuft of flagella and sheath with smooth surface. Bar = 1 µm. (c) Cells with and without a sheath and empty sheaths. Bar = 10 µm. All figures reproduced with permission from Mulder and Deinema (1992).

sources can be assimilated by strains of S. natans, but the substrate utilization patterns of strains within this species differ widely. In contrast to most Leptothrix strains, S. natans is able to assimilate relatively high concentrations of substrates from which it synthesizes considerable amounts of cellular material. Cells may contain large amounts of poly-β-hydroxybutyrate either as numerous small globules or as a few large globules (Fig. 12a). Polysaccharides may also accumulate. The synthesis of both reserve compounds is stimulated by a high carbon/nitrogen ratio in the medium or by oxygen deficiency (Mulder and Van Veen, 1963).

Preservation Stock cultures of S. natans on agar slants of the previously described media can be stored for about 3 months at 4°C. Addition of 2–3 ml of sterile tap water to the agar slants may prolong the viability for another 3 months. Preservation for longer periods is accomplished by common lyophilization techniques; however, it must be stressed that some Sphaerotilus strains do not survive lyophilization. For the long-term preservation of these strains, freezing in liquid nitrogen can be applied.

Physiology Metabolism Sphaerotilus is rarely found associated with deposits of metal oxides in natural

environments and typically thrives in habitats with normal concentrations of ferrous iron. So far, the mechanism of iron oxidation in this microorganism is poorly understood and it could be that in S. natans, iron oxidation is only a side effect of a biological reaction with a different metabolic function. Sphaerotilus natans is obligately aerobic and respiratory, but it can grow well with low concentrations of oxygen. It can readily adapt to various nutrient concentrations, growth temperatures (10–40°C) and pH values (pH 5.4–9.0), but seems to be sensitive to an increase in NaCl concentration (upper limit between 0.3 and 0.7%; Dondero, 1975). Glucose is dissimilated via the phosphogluconoate pathway and the tricarboxylic cycle. In the presence of glucose, the oxidation of other sugars, amino acids, and compounds of the tricarboxylic acid cycle seems to be repressed (Dondero, 1975). Structure and Composition of the Sheath The sheath of S. natans is, like the sheath of Leptothrix, resistant to proteases and composed of a complex of polysaccharide, protein and lipid. Romano and Peloquin (1963) detected in purified sheath material 36% carbohydrate, 28% protein and 5.2% lipid, whereas Takeda et al. (1998) obtained values of 54.1, 12.2 and 1–3%, respectively. These discrepancies may be due to the investigation of different strains of S. natans or the application of various cultivation conditions or sheath purification methods. The surface

CHAPTER 3.2.11

of the S. natans sheath is smooth and covered with an acidic exopolysaccharide composed of fucose, galactose, glucose and glucuronic acid (Gaudy and Wolfe, 1962). Slime production by exopolysaccharide secretion is much more pronounced in this species than in members of the genus Leptothrix. In contrast to the exopolysaccharide of the slime capsule, the sheath carbohydrate is free of acidic sugars and contains only glucose and galactosamine which is probably in its N-acetylated form. A heteropolysaccharide composed of glucose and galactosamine (in a molar ratio of 1 : 4) can be released from the sheath by hydrazine treatment which completely degrades the sheath structure (Takeda et al., 1998). The sheath of S. natans can be also degraded enzymatically by a kind of eliminase (produced by a Paenibacillus sp.) which attacks the polysaccharide moiety of the sheath (Takeda et al., 2000). Although the sheath structure is resistant to protease treatment, part of the sheath protein appears to be sensitive to protease attack and can be removed from the sheath by treatment with agents that reduce disulfide bonds. However, in contrast to the Leptothrix sheath, disulfide bonds apparently do not play a role in maintaining the sheath structure of S. natans. In sheaths treated with protease, the most abundant amino acids were glycine and cysteine. Only three or four other major amino acids were detected, and it was concluded that the sheath protein may consist of repeating subunits of a small peptide which is crosslinked with a polysaccharide backbone (Takeda et al., 1998). The revealed differences in the fine structure and composition of sheaths from Sphaerotilus and Leptothrix may be an indication for different functions of these structures for the respective microorganisms.

Acknowledgments. I am grateful to W. C. Ghiorse for providing unpublished photographs.

Literature Cited Adams, L. F., and W. C. Ghiorse. 1986. Physiology and ultrastructure of Leptothrix discophora SS-1. Arch. Microbiol. 145:126–135. Amann, R. I., W. Ludwig, and K.-H. Schleifer. 1995. Phylogenetic identification and in situ detection of individual microbial cells without cultivation. Microbiol. Rev. 59:143–169. Brouwers, G. J., E. Vijgenboom, P. L. A. M. Corstjens, J. P. M. de Vrind, and E. W. de Vrind-de Jong. 2000. Bacterial Mn2+ oxidizing systems and multicopper oxidases: an overview of mechanisms and functions. Geomicrobiol. J. 17:1–24.

The Genera Leptothrix and Sphaerotilus

775

Carlile, M. J., and W. L. Dudeney. 2001. The discs of Leptothrix: Lost for 89 years. Microbiology 147:1393– 1394. Cataldi, M. S. 1939. Estudio fisió logico y sistem´tico de algunas Chlamydobacteriales (thesis). University of Buenos Aires. Buenos Aires, Argentina. 1–96. Charlet, E., and W. Schwartz. 1954. Beiträge zur Biologie der Eisenmikroben. I: Untersuchungen ü ber die Lebensweise von Leptothrix ochracea und einigen begleitenden Eisenmikroben. Schweiz. Z. Hydrol. 16:318–341. Cholodny, N. 1926. Die Eisenbakterien. Beiträge zu einer Monographie. Pflanzenforsch G. Fischer. Jena, Germany. 4:1–162. Corstjens, P. L. A. M., J. P. M. de Vrind, P. Westbroek, and E. W. de Vrind-de Jong. 1992. Enzymatic iron-oxidation by Leptothrix discophora: identification of an ironoxidizing protein. Appl. Environ. Microbiol. 58:450–454. De Vrind-de Jong, E. W., P. L. A. M. Corstjens, E. S., Kempers, P. Westbroek, and J. P. M. de Vrind. 1990. Oxidation of manganese and iron by Leptothrix discophora: use of N,N,N′,N′-tetramethyl-p-phenylenediamine as an indicator of metal oxidation. Appl. Environ. Microbiol. 56:3458–3462. Dondero, N. C. 1975. The Sphaerotilus-Leptothrix group. Ann. Rev. Microbiol. 3:77–107. Dymond, J., R. W. Collier, and M. E. Watwood. 1989. Bacterial mats from Crater Lake, Oregon and their relationship to possible deep-lake hydrothermal venting. Nature 342:673–675. Eikelboom, D. H. 1975. Filamentous organisms observed in activated sludge. Water Res. 9:365–388. Emerson, D., and W. C. Ghiorse. 1992. Isolation, cultural maintenance, and taxonomy of a sheath-forming strain of Leptothrix discophora and characterization of manganese-oxidizing activity associated with the sheath. Appl. Environ. Microbiol. 58:4001–4010. Emerson, D., and W. C. Ghiorse. 1993a. Role of disulfide bonds in maintaining the structural integrity of the sheath of Leptothrix discophora SP-6. J. Bacteriol. 175:7819–7827. Emerson, D., and W. C. Ghiorse. 1993b. Ultrastructure and chemical composition of the sheath of Leptothrix discophora SP-6. J. Bacteriol. 175:7808–7818. Emerson, D., and N. P. Revsbech. 1994. Investigation of an iron-oxidizing microbial mat community located near Aarhus, Denmark: field studies. Appl. Environ. Microbiol. 60:4022–4031. Esposito, A., F. Pagnanelli, A. Lodi, C. Solisio, and F. Veglio. 2001. Biosorption of heavy metals by Sphaerotilus natans: An equilibrium study at different pH and biomass concentrations. Hydrometallurgy 60:129–141. Felsenstein, J. 1982. Numerical methods for inferring phylogenetic trees. Q. Rev. Biol. 57:379–404. Gaudy, E., and R. S. Wolfe. 1961. Factors affecting filamentous growth of Sphaerotilus natans. Appl. Microbiol. 9:580–584. Gaudy, E., and R. S. Wolfe. 1962. Composition of an extracellular polysaccharide produced by Sphaerotilus natans. Appl. Microbiol. 10:200–205. Ghiorse, W. C., and S. D. Chapnick. 1983. Metal-depositing bacteria and the distribution of manganese and iron in swamp waters. In: R. Hallberg (Ed.) Environmental Biogeochemistry. Publ. House/FRN. Stockholm, Sweden. Ecol. Bull. 35:367–376. Ghiorse, W. C., and H. L. Ehrlich. 1992. Microbial Biomineralization of Iron and Manganese. In: R. W. Fitzpatrick

776

S. Spring

and H. C. W. Skinner (Eds.) Iron and manganese biomineralization processes in modern and ancient environments. Catena. Cremlingen-Destedt, Germany. Catena Supplement 21:75–99. Ghiorse, W. C., D. N. Miller, R. L. Sandoli, and P. L. Siering. 1996. Applications of laser scanning microscopy for analysis of aquatic microhabitats. Microsci. Res. Tech. 33:73–86. Hallbeck, L., F. Stå hl, and K. Pedersen. 1993. Phylogeny and phenotypic characterization of the stalk-forming and iron-oxidizing bacterium Gallionella ferruginea. J. Gen. Microbiol. 139:1531–1535. Kalmbach, S., W. Manz, J. Wecke, and U. Szewzyk. 1999. Aquabacterium gen. nov., with description of Aquabacterium citratiphilum sp. nov., Aquabacterium parvum sp. nov. and Aquabacterium commune sp. nov., three in situ dominant bacterial species from the Berlin drinking water system. Int. J. Syst. Bacteriol. 49:769–777. Kämpfer, P., D. Weltin, D. Hoffmeister, and W. Dott. 1995. Growth requirements of filamentous bacteria isolated from bulking and scumming sludge. Water Res. 29:1585– 1588. Kämpfer, P. 1997. Detection and cultivation of filamentous bacteria from activated sludge. FEMS Microbiol. Ecol. 23:169–181. Kämpfer, P. 1998. Some chemotaxonomic and physiological properties of the genus Sphaerotilus. Syst. Appl. Microbiol. 21:245–250. Kersters, K., and J. De Ley. 1984. Genus Alcaligenes Castellani and Chalmers 1919, 936AL. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology, 1st ed. Williams and Wilkins. Baltimore, MD. 1:361–373. Kü tzing, F. T. 1833. Beitrag zur Kenntnis ü ber die Entstehung und Metamorphose der niederen vegetalischen Organismen, nebst einer systematischen Zusammensetzung der hierher gehörigen niederen Algenformen. Linnaea 8:335–387. Kü tzing, F. T. 1843. Phycologia Generales. Leipzig, Germany. Lieske, R. 1919. Zur Ernährungsphysiologie der Eisenbakterien. Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. II 49:413–425. Ludwig, W., and O. Strunk. 1997. ARB—a software environment for sequence data. (http://www.arb-home.de/pub/ ARB/documentation/arb.ps). Malmqvist, ,ÅT. Welander, E. Moore, A. Ternström, G. Molin, G., and I.-M. Stenström. 1994. Ideonella dechloratans gen. nov., sp. nov., a new bacterium capable of growing anaerobically with chlorate as an electron acceptor. Syst. Appl. Microbiol. 17:58–64. Molisch, H. 1910. Die Eisenbakterien. G. Fischer. Jena, Germany. 1–83. Mulder, E. G., and W. L. van Veen. 1963. Investigations on the Sphaerotilus-Leptothrix group. Ant. v. Leeuwenhoek 29:121–153. Mulder, E. G. 1989a. Genus Leptothrix Kü tzing 1843, 198AL. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology, 1st ed. Williams and Wilkins. Baltimore, MD. 3:1998–2003. Mulder, E. G. 1989b. Genus Sphaerotilus Kü tzing 1833, 386AL. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology, 1st ed. Williams and Wilkins. Baltimore, MD. 3:1994– 1998.

CHAPTER 3.2.11 Mulder, E. G., and M. H. Deinema. 1992. The sheated bacteria. In: A. Balows, H.-G. Trü per, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. Berlin, Germany. 3:2612–2624. Nelson, Y. M., L. W. Lion, M. L. Shuler, and W. C. Ghiorse. 1999. Lead binding to metal oxide and organic phases of natural aquatic biofilms. Limnol. Oceanogr. 44:1715– 1729. Olesen, B. H., R. Avci, and Z. Lewandowski. 2000. Manganese dioxide as a potential cathodic reactant in corrosion of stainless steels. Corrosion Sci. 42:211–227. Palleroni, N. J. 1984. Genus I. Pseudomonas Migula 1894, 237Al. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. C. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology, 1st ed. Williams and Wilkins. Baltimore, MD. 1:141–199. Pellegrin, V., S. Juretschko, M. Wagner, and G. Cottenceau. 1999. Morphological and biochemical properties of a Sphaerotilus sp. Isolated from paper mill slimes. Appl. Environ. Microbiol. 65:156–162. Phaup, J. D. 1968. The biology of Sphaerotilus species. Water Res. 2:597–614. Präve, P. 1957. Untersuchungen ü ber die Stoffwechselphysiologie des Eisenbakteriums Leptothrix ochracea Kü tzing. Arch. Mikrobiol. 27:33–62. Pringsheim, E. G. 1949a. Iron Bacteria. Biol. Rev. Cambridge Philos. Soc. 24:200–245. Pringsheim, E. G. 1949b. The filamentous bacteria Sphaerotilus, Leptothrix, Cladothrix, and their relation to iron and manganese. Philos. Trans. R. Soc. London Ser. B. 233:453–482. Rao, T. S., T. N. Sairam, B. Viswanathan, and K. V. K. Nair. 2000. Carbon steel corrosion by iron oxidising and sulphate reducing bacteria in a freshwater cooling system. Corrosion Sci. 42:1417–1431. Richard, M. G., G. P. Shimizu, and D. Jenkins. 1985. The growth physiology of the filamentous organism Type 021N and its significance to activated sludge bulking. J. Water Pollut. Control Fed. 57:1152–1162. Romano, A. H., and J. P. Peloquin. 1963. Composition of the sheath of Sphaerotilus natans. J. Bacteriol. 86:252–258. Roth, A. W. 1797. Catalecta botanica quibus plantae novae et minus cognitae describuntur atque illistrantur. Lipsiae in Bibliopolio I.G. Mulleriano. fasc. 1. Rouf, M. A., and J. L. Stokes. 1964. Morphology, nutrition, and physiology of Sphaerotilus discophorus. Arch. Mikrobiol. 49:132–149. Saitou, N., and M. Nei. 1987. The neighbor-joining method: a new method for reconstructing phylogenetic trees. Molec. Biol. Evol. 4:406–425. Schwers, H. 1912. Megalothrix discophora, eine neue Eisenbakterie. Zentrbl. Bakteriol. Parasitenkd. Infektkrankh. Hyg. Abt. II 33:273–276. Seviour, E. M., C. Williams, B. DeGrey, J. A. Soddell, R. J. Seviour, and K. C. Lindrea. 1994. Studies on filamentous bacteria from Australian activated sludge plants. Water Res. 28:2335–2342. Siering, P. L., and W. C. Ghiorse. 1997a. Development and application of 16S rRNA-targeted probes for detection of iron- and manganese-oxidizing sheathed bacteria in environmental samples. Appl. Environ. Microbiol. 63:644–651. Siering, P. L., and W. C. Ghiorse. 1997b. PCR detection of a putative manganese oxidation gene (mofA) in environmental samples and assessment of mofA gene homology among diverse manganese-oxidizing bacteria. Geomicrobiol. J. 14:109–125.

CHAPTER 3.2.11 Skerman, V. B. D., V. McGowan, and P. H. A. Sneath. 1980. Approved Lists of Bacterial Names. Int. J. Syst. Bacteriol. 30:225–420. Solisio, C., A. Lodi, A. Converti, and M. Del Borghi. 2000. The effect of acid pre-treatment on the biosorption of chromium(III) by Sphaerotilus natans from industrial wastewater. Water Res. 34:3171–3178. Sommaruga, R., and R. Psenner. 1995. Permanent presence of grazing-resistant bacteria in a hypertrophic lake. Appl. Environ. Microbiol. 61:3457–3459. Spring, S., P. Kämpfer, W. Ludwig, and K.-H. Schleifer. 1996. Polyphasic characterization of the genus Leptothrix: New descriptions of Leptothrix mobilis sp. nov. and Leptothrix discophora sp. nov., nom. rev. and emended description of Leptothrix cholodnii emend. Syst. Appl. Microbiol. 19:634–643. Stein, L. Y., M. T. La Duc, T. J. Grundi, and K. H. Nealson. 2001. Bacterial and archaeal populations associated with freshwater ferromanganous micronodules and sediments. Environ. Microbiol. 3:10–18. Stokes, J. L. 1954. Studies on the filamentous sheathed iron bacterium Sphaerotilus natans. J. Bacteriol. 67:278–291. Stokes, J. L., and M. T. Powers. 1965. Formation of rough and smooth strains of Sphaerotilus discophorus. Ant. v. Leeuwenhoek 31:157–164. Sunda, W. G., and D. J. Kieber. 1994. Oxidation of humic substances by manganese oxides yields low-molecularweight organic substrates. Nature 367:62–64. Suyama, T., T. Shigematsu, S. Takaichi, Y. Nodasaka, S. Fujikawa, H. Hosoya, Y. Tokiwa, T. Kanagawa, and S. Hanada. 1999. Roseateles depolymerans gen. nov., sp. nov., a new bacteriochlorophyll a-containing obligate aerobe belonging to the β-subclass of the Proteobacteria. Int. J. Syst. Bacteriol. 49:449–457. Takeda, M., F. Nakano, T. Nagase, K. Iohara, and J.-I. Koizumi. 1998. Isolation and chemical composition of the sheath of Sphaerotilus natans. Biosci. Biotechnol. Biochem. 62:1138–1143.

The Genera Leptothrix and Sphaerotilus

777

Takeda, M., K. Iohara, S. Shinmaru, I. Suzuki, and J.-I. Koizumi. 2000. Purification and properties of an enzyme capable of degrading the sheath of Sphaerotilus natans. Appl. Environ. Microbiol. 66:4998–5004. Van Veen, W. L., E. G. Mulder, and M. H. Deinema. 1978. The Sphaerotilus-Leptothrix Group of Bacteria. Microbiol. Rev. 42:329–356. Venosa, A. D. 1975. Lysis of Sphaerotilus natans swarm cells by Bdellovibrio bacteriovorus. Appl. Microbiol. 29:702– 705. Wagner, M., R. Amann, P. Kämpfer, B. Assmus, A. Hartmann, P. Hutzler, N. Springer, and K.-H. Schleifer. 1994. Identification and in situ detection of Gram-negative filamentous bacteria in activated sludge. Syst. Appl. Microbiol. 17:405–417. Wen, A., M. Fegan, C. Hayward, S. Chakraborty, and L. I. Sly. 1999. Phylogenetic relationsships among members of the Comamonadaceae, and description of Delftia acidovorans (den Dooren de Jong 1926 and Tamaoka et al 1987) gen. nov., comb. nov. Int. J. Syst. Bacteriol. 49:567– 576. Williams, M. W., and R. F. Unz. 1985. Isolation and characterization of filamentous bacteria present in bulking activated sludge. Appl. Microbiol. Biotechnol. 22:273–280. Winogradsky, S. 1888. Ü ber Eisenbakterien. Bot. Zeitschr. 46:261–270. Winogradsky, S. 1922. Eisenbakterien als Anorgoxydanten. Zentralbl. Bakteriol. Parasitenkd. Infektionskr. Hyg. Abt. II 57:1–21. Winston, V., and T. L. Thompson. 1979. Isolation and characterization of a bacteriophage specific for Sphaerotilus natans which contains an unusual base in its deoxyribonucleic acid. Appl. Environ. Microbiol. 37: 1025–1030. Ziegler, M., M. Lange, and W. Dott. 1990. Isolation and morphological and cytological characterization of filamentous bacteria from bulking sludge. Water Res. 24:1437– 1451.

Prokaryotes (2006) 5:778–811 DOI: 10.1007/0-387-30745-1_36

CHAPTER 3.2.12 ehT

c i hpor totuaoht iL

gn i z i d i xO- a i nom A

a i re t caB

The Lithoautotrophic Ammonia-Oxidizing Bacteria HANS-PETER KOOPS, ULRIKE PURKHOLD, ANDREAS POMMERENING-RÖ SER, GABRIELE TIMMERMANN AND MICHAEL WAGNER

Introduction The lithoautotrophic ammonia-oxidizing bacteria (AOB) are well defined by their fundamental metabolism. Ammonia serves as the sole energy source and carbon dioxide is used to fulfill the carbon need. Together with the lithotrophic nitrite-oxidizing bacteria (see the chapter Nitrite Oxidizing Bacteria in this Volume), the AOB catalyze the so-called “nitrification process” (NH3 → NO2 − → NO3 − ), which has a key position in natural nitrogen cycling. Details of the biochemistry of the AOB are reported in the chapter on Oxidation of Inorganic Nitrogen Compounds as an Energy Source in Volume 2. This chapter focuses on the phylogeny, taxonomy, and diversity of AOB as well as on their distribution in nature. Winogradsky in his fundamental investigations on AOB already postulated the existence of a great diversity of species within this guild (Winogradsky, 1892; Winogradsky and Winogradsky, 1937; Winogradsky and Winogradsky, 1933). However, isolation of AOB turned out to be difficult and was neglected for several decades after Winogradsky’s classical studies. Therefore, Nitrosomonas europaea was the only species in culture for a long period. Beginning in the 1960s, S. W. Watson from the Woods Hole Research Center (Woods Hole, MA, United States) and others started a new era of isolation and culturing of novel species of AOB. Since that time, 16 species of AOB have been described and named (Watson, 1965; Watson et al., 1971c; Harms et al., 1976; Jones et al., 1988; Koops et al., 1990; Koops et al., 1991), and the existence of many other cultured species has been reported in the literature (Koops and Harms, 1985; Stehr et al., 1995a). Meanwhile, the phylogeny of the described species and the majority of the yet unnamed cultured species has become well established (Woese et al., 1984; Woese et al., 1985; Head et al., 1993; Teske et al., 1994; Pommerening-Röser et al., 1996; Purkhold et al., 2000; Purkhold et al., 2003; see the section on Phylogeny of AOB in this Chapter), providing a sufficient basis for investigations of natural AOB

populations employing classical as well as molecular techniques. In particular, the polymerase chain reaction (PCR)-assisted retrieval of 16S rRNA gene or amoA gene (encoding the active site subunit of the AOB key enzyme ammonia monooxygenase) sequences from the environment (e.g., Stephen et al., 1996; Purkhold et al., 2000) and the direct identification of AOB via fluorescence in situ hybridization FISH (e.g., Juretschko et al., 1998) allow the diversity of AOB to be monitored and their abundance in situ to be quantified. Results obtained from such investigations together with results from laboratory studies of cultured species have led to a better understanding of the ecology of this bacterial guild. The following sections describe enrichment and isolation techniques of AOB (see the section on Enrichment, Isolation, and Maintenance in this Chapter), summarize the present knowledge on their taxonomy (see the section on General Characteristics of the Genera and Species of AOB in this Chapter) and phylogeny (see the section on Phylogeny of AOB in this Chapter), and provides an overview of available techniques for investigating AOB (see the “Methods Useful for In Situ Detection of AOB” in this Chapter). The last section aims to reveal possible correlation between ecophysiological characteristics and distribution patterns of the distinct species or groups of species of AOB (see the section on Dominant Populations of AOB in Different Environments in this Chapter).

Enrichment, Isolation, and Maintenance Isolation procedures in general are targeted to the numerically most abundant AOB of the respective sample site. Therefore, the first step of enrichment should be a most probable number (MPN) dilution series in media, as far as possible reflecting the conditions of the environment under investigation. For example, in enrichments from acidic environments, low pH of the medium and addition of urea as the sole substrate for

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

energy generation is useful to isolate members of the dominant subpopulation of AOB. In general, the most important factors are the ammonia and the salt concentration in the medium. In some cases, temperature also may be of importance. Two alternative second steps on the way to isolation have been most successful. One option is additional series of dilutions, always using the highest positive dilution of the foregoing MPN series as inoculum. The other option is plating of the bacteria from the highest positive dilution of the original MPN dilution series. It is important to keep in mind that AOB sometimes grow extremely slowly. Therefore, a positive reaction in the highest dilution of an MPN series sometimes may become visible (via pH indicator) not before incubation for two or three weeks, and colony picking from agar plates should await a minimum growth period of 6–8 weeks. Standard media, useful for enrichment cultures or pure cultures, are presented in Tables 1A and 1B. For stock cultures, calcium carbonate (5 g per liter) or N-(2-hydroxyethyl)piperazine-N′2-ethanesulfonic acid (HEPES; 4.77 g per liter) can be used as a pH buffer. In working cultures, the pH should be adjusted by addition of a 10%

solution of sodium hydrogen carbonate. This addition is either performed manually (with the aid of a pH indicator) or by using an automatic pH controller. Since ammonia and not ammonium is the substrate of AOB, the pH optimum in part depends on the concentration of the added ammonium salt. Ten mM ammonium salts at a pH of about 8.0 can be regarded as good standard for sufficient growth of AOB cultures. Large volume cultures should be stirred to guarantee a sufficient supply with oxygen. In general, 30°C is a suitable growth temperature. Stock cultures should be transferred to fresh medium every 3–4 months. Storage of AOB cells in liquid nitrogen is possible, whereas freezedrying of AOB cells in general has not been successful.

General Characteristics of the Genera and Species of AOB The taxonomic framework of the AOB stems from early investigations of Winogradsky (Winogradsky, 1892; Migula, 1900; Buchanan, 1917; Winogradsky and Winogradsky, 1933). The

Table 1A. Different growth media for lithotrophic ammonia oxidizers. Medium no.b Ingredient Distilled water (ml) Seawater (ml) NH4Cl (mg) (NH4)2SO4 (mg) MgSO4 × 7H2O (mg) CaCl2 × 2H2O (mg) KH2PO4 (mg) K2HPO4 (mg) KCl (mg) NaCl (mg) Chelated iron (mg) FeSO4 × 7H2O (µg) Fe-EDTA (mg) Na2MoO4 × 2H2O (µg) (NH4)6Mo7O24 × 4H2O (µg) MnCl2 × 4H2O (µg) MnSO4 × 4H2O (µg) CoCl2 × 6H2O (µg) CuSO4 × 5H2O (µg) ZnSO4 × 7H2O (µg) H3BO3 (µg) Phenol red, 0.5% (ml) Cresol red, 0.05% (ml) a

1 1,000

2 1,000

3

130.0 200.0 20.0

4

5

1,000 1,320.0

600 400 500.0

1,000 535.0

500.0 40.0 40.0 200.0

49.3 147.0 54.4

87.0

200.0 20.0 50.0 114.0

74.4 584.0 1.0

1.0 973.1

0.5 100.0

1.0 37.1

200.0

2.0 44.6

2.0 20.0 100.0 0.1

25.0 43.1 49.4

1.0

2.0 20.0 100.0 1.0

1.0

1.0

Suitable buffers are 5.0g/l CaCO3 and 4g/l HEPES, respectively. Medium no. 1 is from Soriano and Walker (1968), for terrestrial strains. Medium no. 2 is from Watson (1971b), for terrestrial strains. Medium no. 3 is from Krü mmel and Harms (1982), for terrestrial strains. Medium no. 3 is from Watson (1965), for marine strains. Medium no. 5 is from Koops et al. (1976), for brackish-water strains.

b

779

780

H.-P. Koops et al.

CHAPTER 3.2.12

isolates originally were categorized into genera on the basis of the shape of cells. Later, the arrangement of their intracytoplasmic membranes was introduced as a second basic character. Using these criteria, the genera Nitrosomonas, Nitrosococcus, Nitrosospira, Nitrosolobus and Nitrosovibrio have been established (Watson et al., 1981; Watson et al., 1989; Koops and Möller, 1992). In the following, a brief listing of the distinguishing morphological features of the five recognized genera of AOB is given and the distinct csultured species are described. Differential characteristics of the genera of AOB and of the species of AOB are presented in Tables 2 and 3, respectively.

The Genera Genus NITROSOMONAS (Winogradsky, 1892) Cells generally are rod shaped, but sometimes spherical. Extensive intracytoplasmic membranes are arranged as peripherally located flattened vesicles. Sometimes intrusions of membranes into the protoplasm are observed (Fig. 1). Genus NITROSOSPIRA (Winogradsky and Winogradsky, 1933) Cells are tightly closed spirals and occasionally vibrio shaped. Extensive intracytoplasmic membranes are missing. Intru-

Table 1B. Different media used to ascertain that cultures of ammonia oxidizing bacteria are free of heterotrophic contaminants. Medium no.b Ingredient Distilled water Seawater Yeast extract (Difco) Peptone (Difco) Beef extract (Difco)

1

2

3

1,000ml

600ml 400ml 0.5g 0.5g 0.5g

250ml 750ml 0.5g 0.5g 0.5g

0.5g 0.5g 0.5g

a

The pH of these media should be adjusted to 7.3. Media no. 1, 2 and 3 are from Koops et al. (1976). Medium no. 1 is for terrestrial and freshwater strains, respectively; no. 2 for brackish-water strains; and no. 4 for marine strains.

b

sions of membranes into the protoplasm are sporadically observed (Fig. 2). Genus NITROSOVIBRIO (Harms et al., 1976) Cells are vibrio shaped. Extensive intracytoplasmic membranes are missing. Intrusions of membranes into the protoplasm are documented (Fig. 3). Genus NITROSOLOBUS (Watson et al., 1971c) Cells are pleomorphic lobes compartmentalized by the cytoplasmic membrane (Fig. 4). Genus NITROSOCOCCUS (Winogradsky, 1892) This genus solely represents the gamma-proteobacterial AOB. Both described species, N. oceani and N. halophilus, are characterized by large spherical to ellipsoidal cells revealing extensive intracytoplasmic membranes, arranged as a central stack of vesicles (Fig. 5).

The Cultured Species Since all AOB possess the same fundamental metabolism and morphological differences are per se limited within this group, identification on the species level often is difficult if exclusively phenotypic characters are used. The three genera Nitrosolobus, Nitrosovibrio and Nitrosospira, which phylogenetically form the Nitrosospira lineage (see the section on Phylogeny of AOB in this Chapter), are severely affected by this limitation. Nitrosomonas species within one of the distinct lineages of this genus often cannot be unambiguously identified using phenotypic characters, while species affiliated with different lineages can more easily be distinguished. In general, DNA-DNA hybridization is the method of choice to define closely related species. The above-mentioned phenotypic similarities among closely related AOB has hampered in many cases formal description of genotypically well-defined, cultured species. Below, important features of AOB species are listed.

Table 2. Characteristics of the genera of the ammonia-oxidizing bacteria. Characteristics Cell shape Cell size (µm) Flagellation of motile cells Arrangement of intracytoplasmic membranes

Nitrosococcus

Nitrosolobus

Spherical to ellipsoidal 1.5–1.8 × 1.7–2.5 Tuft of flagella

Pleomorphic lobate

Straight rods

Nitrosomonas

1.0–1.5 × 1.0–2.5 Peritrichous

Central stacks of vesicles

Compartmentalizing membranes

0.7–1.5 × 1.0–2.4 Polar to subpolar Peripheral flattened vesicles

Nitrosospira

Nitrosovibrio

Tightly coiled spirals 0.3–0.8 × 1.0–8.0 Peritrichous

Slender curved rods 0.3–0.4 × 1.1–3.0 Polar to subpolar Invaginations

Invaginations

50.6–51.4 47.9–48.5 53.8 49.3 45.6–46.0 47.9 45.6–46.0 49.4–50.0 47.4–48.0 45.7–46.3 45.5–46.1 53.5 53.9 54 50–51 50–51

G+C (mol%) − − − − − + + + + + + +/− +/− +/− + −

Carboxysomes − + + − − + − − − − − ND ND ND ND ND 42–59 ND ND ND ND ND

50–52

14–43 19–46 1.9–4.2

30–61

400 600 400 250 250 100 200 50 200 400 400 50 100 200 1000 500

Maximum ammonia tolerance NH4Cl (in mN; pH 8.0)

Symbols and Abbreviations: +, present; −, not present; +/−, present in some strains; and ND, no data.

Nitrosomonas europaea Nitrosomonas eutropha Nitrosomonas halophila Nitrosococcus mobilis Nitrosomonas communis Nitrosomonas nitrosa Nitrosomonas ureae Nitrosomonas oligotropha Nitrosomonas marina Nitrosomonas aestuarii Nitrosomonas cryotolerans Nitrosolobus multiformis Nitrosovibrio tenuis Nitrosospira briensis Nitrosococcus oceani Nitrosococcus halophilus

Species

Urease activity

Substrate (NH3) affinity (K, in µM)

Table 3. Characteristics and preferred habitats of described species of the ammonia-oxidizing bacteria.

− − + + − − − − + + + − − − + +

Salt requirement 400 400 900 500 250 300 200 150 800 600 550 200 100 250 1100 1800

Maximum salt tolerance (in mM)

Marine environments Soils (not acid) Soils, rocks and freshwater Soils, rocks and freshwater Marine environments Marine environments and salt lakes

Marine environments

Oligotrophic freshwater and natural soils

Soils (not acid) eutrophic freshwater

Sewage disposal plants, eutrophic freshwater and brackish water

Preferred habitats

CHAPTER 3.2.12 The Lithoautotrophic Ammonia-Oxidizing Bacteria 781

782

H.-P. Koops et al.

CHAPTER 3.2.12

Fig. 1A

Fig. 1B 5 mm

5 mm

0.2 mm

C

IM c

IM

0.3 mm

Fig. 1. The genus Nitrosomonas. Phase contrast photomicrographs (A) and electron micrographs of thin sections (B) of cells of different Nitrosomonas species showing the variability of shapes and sizes and the details of their ultrastructure (intracytoplasmic membranes [IM] and carboxysomes [C]).

1 mm

1 mm

3A

2A 0.5 mm

2B 3B Fig. 2. The genus Nitrosospira. Scanning electron micrograph (A) and electron micrograph of thin sections (B) of cells of Nitrosospira species.

0.3 mm

Fig. 3. The genus Nitrosovibrio. Scanning electron micrograph (A) and electron micrograph of thin sections (B) of cells of Nitrosovibrio species.

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

The Species of the Betaproteobacterial AOB The Nitrosomonas Group The cultured species of this genus can be assigned to six distinct, phylogenetically definable lineages (see the section on Phylogeny of AOB in this Chapter), comprising 11 described species. Several further genospecies are in culture but cannot sufficiently be discriminated by phenotypic properties (e.g., Koops and Harms, 1985; Stehr et al., 1995a). In

1 mm

4A

4B

0.5 mm

Fig. 4. The genus Nitrosolobus. Scanning electron micrograph (A) and electron micrograph of thin sections (B) of cells of Nitrosolobus species.

general, described species belonging to different lineages of Nitrosomonas are phenotypically well distinguishable (Table 3). However, within the distinct Nitrosomonas lineages, species discrimination in many cases is as impossible as observed for the Nitrosospira lineage. The Nitrosomonas europaea/Nc. mobilis Lineage This lineage comprises four described species, all being characterized by a relatively high salt tolerance. Two of these species, N. halophila and Nc. mobilis, have an obligatory but moderate salt requirement. All reveal relatively high affinity constants for ammonia (50– 100 µM), and all cultured strains are urease negative. Consistent with these findings the members of this group seem to prefer eutrophicated aquatic environments with heightened ionic strength. The levels of phylogenetic interrelationships among the four species of this lineage are different. While N. europaea and N. eutropha reveal absolute close relationship with each other (15–16% DNA-DNA similarity), they show only minimal DNA-DNA similarity values with N. halophila, and DNA-DNA similarities are even missing with Nc. mobilis, as measured with the S1 nuclease technique (PommereningRöser et al., 1996). Nitrosomonas europaea (Winogradsky, 1890; Watson, 1971a) Cells (0.8–1.1 × 1.0–1.7 µm) are short rods with pointed ends. Motility is not observed, and carboxysomes are missing. Often found in sewage disposal plants, cells are occasionally observed in eutrophicated freshwaters or fertilized soils. They have no obligate salt requirement, but do have a striking tolerance for

Fig. 5A Fig. 5B 0.5 mm

Fig. 5. The genus Nitrosococcus. Phase-contrast photomicrographs of whole cells (A) and electron micrographs of thin sections of cells (B) of Nitrosococcus halophilus. (C) Electron micrograph of a Nitrosococcus halophilus cell shadowed with chromium showing a tuft of flagella. (D) Freeze-etch electron micrograph of Nitrosococcus oceani showing the macromolecular arrangement of the two layers outside of the envelope.

783

5 mm

Fig. 5C Fig. 5D

1 mm

0.1 mm

784

H.-P. Koops et al.

CHAPTER 3.2.12

increasing salt concentration up to 400 mM NaCl. The G+C content of DNA is 50.6–51.4 mol%.

preferred habitat and freshwater only an occasional habitat. The G+C content of DNA is 45.6– 46.0 mol%.

Nitrosomonas eutropha (Koops et al., 1991) Cells occur singly or as short chains and are pleomorphic, rod- to pear-shaped, with one or both ends are pointed. Cells (1.0–1.3 × 1.6–2.3 µm) are motile, and carboxysomes are observed. The species is commonly distributed in sewage disposal plants, and occasionally it has been isolated from other eutrophicated environments. The species has no salt requirement but is tolerant to high salt concentration up to 400 mM NaCl. The G+C content of DNA is 47.9–48.5 mol%.

Nitrosomonas nitrosa (Koops et al., 1991) Cells (1.3–1.5 × 1.4–2.2 µm) are spheres or rods with rounded ends. Motility is not observed. Carboxysomes are present. Although eutrophicated freshwaters seem to be the preferred habitats, strains occasionally have been isolated from marine environments and a wastewater treatment plant. The G+C content of the DNA is 47.9 mol%.

Nitrosomonas halophila (Koops et al., 1991) Cells (1.1–1.5 × 1.5–2.2 µm) occasionally appear coccoid. Motile cells possess a tuft of flagella, and carboxysomes are observed. Cells have an obligate salt requirement, and some isolates are alkali tolerant up to pH 10. Cells tolerate high salt concentration (up to 900 mM NaCl). Strains were isolated from the North Sea and from soda lakes located in Mongolia (Sorokin et al., 2001). The G+C content of DNA is 53.8 mol%. Nitrosococcus (Nitrosomonas) mobilis (Koops et al., 1976) Cells (1.5–1.7 × 1.5–2.1 µm) are coccoid or rod shaped. Carboxysomes have not been observed. Motile cells possess a tuft of flagella. Cells have an obligate salt requirement. Isolates originate from the North Sea (Koops and Harms, 1985) and from an industrial wastewater treatment plant (Juretschko et al., 1998). The species seems to prefer eutrophic, aquatic environments. The G+C content of DNA is 49.3 mol%.

The Nitrosomonas oligotropha Lineage Two described species and several undescribed genospecies (e.g., Nitrosomonas sp. Nm47, Nitrosomonas sp. Nm86, Nitrosomonas sp. Nm84, and Nitrosomonas sp. Nm59) belonging to this lineage exist in culture (see the section on Phylogeny of AOB in this Chapter). The absolute majority of strains originate from oligotrophic freshwaters, and only occasional isolates are from natural, often moderately acidic (pH about 6.0) soils. All strains investigated thus far reveal strikingly low affinity constants for ammonia (few µM) and almost all isolates are urease positive. All studied strains turned out to be salt sensitive. Nitrosomonas oligotropha (Koops et al., 1991) Cells (0.8–1.2 × 1.1–2.4 µm) are rod shaped or spherical with rounded ends. Cell aggregates, obviously caused by extensive production of exopolymeric materials, are observed in the environment as well as in pure culture. Carboxysomes are missing, motility is not observed. The G+C content of DNA is 49.4–50.0 mol%.

The Nitrosomonas communis Lineage According to phenotypic properties, this lineage is divided into two sublineages. This division is also supported by DNA-DNA similarity data (Pommerening-Röser et al., 1996). One subgroup contains N. communis together with two undescribed cultured genospecies (see the section on Phylogeny of AOB in this Chapter) that are phenotypically not well distinguishable. These three species are urease negative and prefer agricultural soils with neutral pH. The other subgroup is represented by a single species, N. nitrosa. In contrast to the first subgroup, all isolates of N. nitrosa are urease positive, and aquatic environments are the preferred habitats.

The Nitrosomonas marina Lineage This lineage is represented by two described species and one undescribed genospecies (represented by the isolates Nm51 and Nm63; see the section on Phylogeny of AOB in this Chapter). All isolates originate from marine environments and reveal an obligate salt requirement. In general, isolates are urease positive.

Nitrosomonas communis (Koops et al., 1991) Cells (1.0–1.4 × 1.7–2.2 µm) are relatively large rods with rounded ends. Motility is not observed, and carboxysomes are missing. Moderately eutrophicated, pH neutral soils seem to be the

Nitrosomonas marina (Koops et al., 1991) Cells (0.7–0.9 × 1.7–2.2 µm) are slender rods with rounded ends. Motility is not observed, and carboxysomes are missing. The G+C content of DNA is 47.4–48.0 mol%.

Nitrosomonas ureae (Koops et al., 1991) Phenotypically very similar to N. oligotropha but distinguishable by a lower G+C content of the DNA (45.6–46.0 mol%).

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

Nitrosomonas aestuarii (Koops et al., 1991) Phenotypically, this species is very similar to N. marina. Cells (1.0–1.3 × 1.4–2.0 µm) are rod shaped. The G+C content of DNA is 45.7–46.3 mol%. The Nitrosomonas cryotolerans Lineage Only one species has yet been described, and only one strain is available in culture. The only cultured strain of Nitrosomonas cryotolerans (Jones et al., 1988) originates from surface water of the Kasitsna Bay (Alaska). The rod shaped cells (2.0–4.0 × 1.2–2.2 µm) requires salt and is urease positive. Carboxysomes are missing. Motility has not been observed. Intracytoplasmic membranes are not exclusively peripherally arranged, as is typical for the other members of the genus Nitrosomonas, but sometimes membranes intrude deep into the cytoplasm and often surround the nucleoplasm. Cultures can grow at temperatures as low as –5°C. The G+C content of DNA is 45.5–46.1 mol%. The Nitrosomonas sp. Nm143 Lineage In total four isolates belonging to this lineage are available in culture (Ward, 1982; Ward and Carlucci, 1985; Purkhold et al., 2003; see the section on Phylogeny of AOB in this Chapter) but were not described by name. All isolates were obtained from marine systems. The Nitrosospira Lineage Within this group of microorganisms, the three genera Nitrosospira, Nitrosolobus, Nitrosovibrio (each encompassing a single described species) and several genospecies (see the section on Phylogeny of AOB in this Chapter) have been described. While the genera show discriminative morphological characters, phenotypic species discrimination within the distinct genera is very difficult. However, the DNA G+C content differs significantly among genospecies or groups of genospecies (Koops and Harms, 1985). Different affinity constants for the binding of ammonia and different tolerances for increasing ammonia or salt concentrations could eventually serve as distinguishing features. Furthermore, temperature was observed as a factor that could be used to discriminate among species (Jiang and Bakken, 1999). However, no robust phenotypically based system for discriminating between species of the distinct genera is currently available. Unfortunately, this situation is consistent with the lack of sufficient resolution within 16S rRNA genebased phylogenetic trees of the Nitrosospira lineage (see the section on Phylogeny of AOB in this Chapter). THE GENUS NITROSOSPIRA This genus contains many cultured genospecies (see the section on

785

Phylogeny of AOB in this Chapter) that cannot sufficiently be discriminated by phenotypic characteristics (Koops and Harms, 1985). However, two groups, containing two and three distinct species, respectively, can be distinguished on the basis of DNA G+C values around 53.5 and 55.2 mol%, respectively. The existence of these two groups later has been confirmed by DNADNA hybridizations applying the S1 nuclease technique (Pommerening-Röser, 1993). Unfortunately, only a few of the numerous cultured species have yet been included in such investigations. All cultured species are more or less pleomorphic, varying in shape between tightly closed spirals and vibrio-forms. All species, thus far studied, have urease-positive as well as -negative strains at similar frequencies. Species are commonly distributed in natural soils and, occasionally, occur in freshwater environments. Via molecular studies, representatives of Nitrosospira have also been detected in marine environments (McCaig et al., 1994; Stephen et al., 1996; Phillips et al., 1999; Bano and Hollibaugh, 2000; Horz et al., 2000; Hollibaugh et al., 2002; Nicolaisen and Ramsing, 2002). However, marine isolates do not exist and salt requirement has not yet been observed among cultured Nitrosospira strains. Cells of Nitrosospira briensis (Winogradsky and Winogradsky, 1933) are motile cells and possess 1–6 peritrichous flagella. Strains of this species have been isolated from soils. The type strain is urease negative, but other isolates of this species are urease positive. The G+C content of DNA is about 54 mol%. The Genus NITROSOVIBRIO Only one species is in culture. All strains of this species have exclusively vibrio-shaped cells. Pleomorphy, as observed with several genospecies of the genus Nitrosospira, has not yet been detected. The three cultured strains of Nitrosovibrio tenuis (Harms et al., 1976) were isolated from natural soils and are urease positive. Cells are slender curved rods (0.3–0.4 × 1.1–3.0 µm). Motile cells possess 1–4 subpolar flagella. The G+C content of DNA is 53.9 mol%. The Genus NITROSOLOBUS The genus is characterized by the pleomorphic, lobate cells which are compartmentalized by the cytoplasmic membrane. One described species and one genospecies (Nitrosolobus sp. NL5) are available. Both species are distinguishable by cell size and by a significantly different DNA G+C content of 53.5 and 56.5%, respectively (Koops and Harms, 1985). Most of the isolates of both species originate from soils, generally being pH-neutral and agriculturally used. One strain of Nitrosolobus sp. was obtained from a sewage disposal plant.

786

H.-P. Koops et al.

The cells of Nitrosolobus multiformis (1.0–1.5 × 1.0–2.5 µm; Watson et al., 1971c) are pleomorphic, motile, and possess 1–20 peritrichous flagella. Cell division is by constriction. The G+C content of DNA is about 53.5 mol%.

The Species of the Gammaproteobacterial AOB The gammaproteobacterial AOB are represented by only two species of the genus Nitrosococcus: N. oceani and N. halophilus. Both species have a similar morphology. However, both are different in salt requirement, salt tolerance, and ammonia tolerance (Table 3). Moreover, the observation that all cultured strains of N. oceani possess urease whereas the two isolates of N. halophilus are urease negative indicates another discriminating character. However, more strains must be studied to verify these distinguishing features. A third species, N. nitrosus, is no longer available in culture. NITROSOCOCCUS NITROSUS (Migula, 1900) This species is no longer available in culture, and nothing is known about its ecophysiology or its phylogenetic position. This species should be placed on the list of rejected names. NITROSOCOCCUS OCEANI (Watson, 1965; Watson, 1971a; Trüper and de Clari, 1997) This species originally was described as Nitrosocystis oceanus (Watson, 1965) and later renamed “Nitrosococcus oceanus” (Watson, 1971a), before the presently accepted name Nitrosococcus oceani was established (Trü per and de Clari, 1997). Spherical or ellipsoidal cells (1.8–2.2 µm in diameter) occur singly or as pairs. Motile cells possess 1–20 flagella (tuft). Two surface layers exist, being composed of subunits arranged in rectilinear and hexagonal arrays (Fig. 5). Intracytoplasmic membranes are arranged as a stack of parallel, flattened vesicles (Murray and Watson, 1965; Watson, 1965; Watson and Remsen, 1970). All cultured strains originate from marine environments, and all are urease positive. The G+C content of DNA is between 50 and 51 mol%. NITROSOCOCCUS HALOPHILUS (Koops et al., 1990) Both species are morphologically not distinguishable from N. oceani, but differ in salt requirement and salt tolerance as well as in ammonia tolerance (Table 3). The two cultured strains were isolated from an inland salt lake and from a salt lagoon in the Mediterranean Sea, respectively. In contrast to the cultured strains of N. oceani, the two isolates of N. halophilus are urease negative. The G+C content of DNA is 50– 51 mol%.

CHAPTER 3.2.12

Phylogeny of AOB The phylogenetic framework of AOB has been established by comparative 16S rRNA gene sequence analysis of the cultured species. During the last years, this system has been significantly extended by including environmentally retrieved 16S rRNA gene sequences.

The Cultured AOB The first phylogenetic analyses of AOB were carried out by Woese and co-workers in the 1980s (Woese et al., 1984; Woese et al., 1985) and demonstrated that two phylogenetically distinct groups of AOB exist. The major group, containing the genera Nitrosomonas, Nitrosospira, Nitrosovibrio and Nitrosolobus, belongs to the class Betaproteobacteria, while the second group of AOB, represented by two species of the genus Nitrosococcus, is affiliated with the class Gammaproteobacteria (see the chapter Oxidation of Inorganic Nitrogen Compounds as an Energy Source in Volume 2). These results, obtained from 16S rRNA oligonucleotide catalogues, were later confirmed and extended via comparative 16S rRNA gene sequence analysis (Head et al., 1993; Teske et al., 1994; Utaaker et al., 1995; Purkhold et al., 2000). Today, almost complete 16S rRNA gene sequences are available for the 14 described species of AOB affiliated with the Betaproteobacteria. Furthermore, more than 100 16S-rRNA gene sequences of other AOB isolates belonging to this class have been determined. Within the Betaproteobacteria, the AOB form a well-supported monophyletic evolutionary group (Head et al., 1993; Teske et al., 1994). The minimal 16S rRNA gene sequence similarity between recognized members of this group is 89.4% (Nitrosococcus mobilis/Nitrosomonas nitrosa). Figure 6 shows a phylogenetic 16S rRNAbased tree of those AOB (15 nitrosospiras and 19 nitrosomonads) demonstrated to represent different genospecies (DNA-DNA similarity less than 60% and/or 16S rRNA sequence similarity less than 97.5%). The genera Nitrosospira, Nitrosolobus and Nitrosovibrio are closely related, and 16S rRNA phylogeny provides no convincing support for subdivision of this lineage into three genera, although morphological and ecophysiological differences exist between the different genera (see the section on General Characteristics of the Genera and Species of AOB in this Chapter). It has therefore been suggested these genera be lumped into the single genus Nitrosospira (Head et al., 1993). In contrast, the cultured nitrosomonads can be subdivided into six lineages which are consistently

CHAPTER 3.2.12 Fig. 6. 16S rRNA-based phylogenetic tree of the betaproteobacterial AOB. The tree includes only those AOB which have been demonstrated to represent different genospecies (DNADNA similarity < 60%) and for which 16S rRNA gene sequences longer than 1000 nucleotides are available. Strains with DNA-DNA similarity > 60% with each other are given in parenthesis after the respective species name. Described species are depicted in bold. Maximum likelihood, maximum parsimony, and neighbor-joining trees were calculated and merged. Multifurcations connect branches for which a relative order cannot be unambiguously determined by applying different treeing methods. Filled and empty dots indicate parsimony bootstrap values (100 resamplings) above 90% and 70%, respectively. Scale bar represents 10% estimated sequence divergence.

The Lithoautotrophic Ammonia-Oxidizing Bacteria

787

Nitrosomonas europaea Nm50 (Nm103)

Fig. 6

Nitrosomonas eutropha Nm57 Nitrosomonas halophila Nm1 (AN1-5) Nitrosococcus mobilis Nc2 (Nm104, Nm107) Nitrosomonas sp. Nm41 (Nm58) Nitrosomonas sp. Nm33 Nitrosomonas communis Nm2 Nitrosomonas nitrosa Nm90 (Nm148) Nitrosomonas sp. Nm51 (Nm63) Nitrosomonas marina Nm22 Nitrosomonas aestuarii Nm36 Nitrosomonas sp. Nm47 Nitrosomonas oligotropha Nm45 Nitrosomonas ureae Nm10 Nitrosomonas sp. Nm86 Nitrosomonas sp. Nm84 Nitrosomonas sp. Nm59 Nitrosomonas cryotolerans Nm55 Nitrosomonas sp. Nm143 Nitrosospira sp. Nsp1 (Nsp4, Nsp40, F3) Nitrosospira sp. Nsp41 Nitrosospira sp. Nv6 (Nv12) Nitrosospira briensis Nsp10 Nitrosospira sp. Nsp62 Nitrosospira sp. L115 (A4. AF) Nitrosovibrio (Nitrosospira) tenuis Nv1 Nitrosolobus (Nitrosospira) sp. NL5 Nitrosospira sp. Nsp2 Nitrosospira sp. Nsp17 Nitrosolobus (Nitrosospira) multiformis NL13 Nitrosospira sp. Nsp57 (Nsp58) Nitrosospira sp. III7 (B6) Nitrosospira sp. Nsp12 (Nsp5, III2, 40KI) Nitrosospira sp. Nsp65

retrieved using different treeing methods and which (if consisting of more than one sequence) have parsimony bootstrap support of above 90%. This phylogenetic substructure is also retrieved if all betaproteobacterial AOB isolates (56 nitrosomonads and 48 nitrosospiras), for which a 16S rRNA sequence longer than 1000 bases has been determined, are included in the treeing analysis (Figs. 7 and 8). For the different groups within the nitrosomonads, different nomenclature systems are used in the literature (Pommerening-Röser et al., 1996; Stephen, 1996; Purkhold et al., 2000; Aakra et al., 2001b; see also Fig. 7). To avoid confusion, we herein propose a hierarchical system for subdivision of the betaproteobacterial AOB as displayed below for further use. New lineages should only be postulated if 1) almost full-length 16S rRNA sequences for the respective AOB exist and 2) the lineages can be retrieved with all treeing methods with high bootstrap support. The lineages are subdivided into recognized species and genospecies with each of these units possibly

0.10

containing several strains. In the following, a brief list (consistent with the above rules) of those units within the betaproteobacterial AOB is presented: Betaproteobacterial AOB Nitrosospira lineage (Genus Nitrosospira) (Genus Nitrosolobus) (Genus Nitrosovibrio) Nitrosomonas group N. europaea/Nc. mobilis lineage N. communis lineage N. marina lineage N. oligotropha lineage N. cryotolerans lineage Nitrosomonas sp. Nm143 lineage Although 16S rRNA phylogeny of the nitrosospiras does not reveal an obvious substructure, Stephen et al. (1996) suggested subdivision of this group into three “clusters” (clusters 2, 3 and 4, Fig. 8; cluster 1 contains no cultured species,

788

H.-P. Koops et al.

Nitrosomonas sp. TK794 Nitrosomonas sp. IWT202 Nitrosomonas sp. F6 Nitrosomonas eutropha Nm57 Nitrosomonas sp. CNS326 Nitrosomonas sp. GH22 Nitrosomonas sp. G1 Nitrosomonas sp. K1 Nitrosomonas sp. F5

Fig. 7

N. europaeal Nc. mobilis lineage (Cluster 7) 92.4

CHAPTER 3.2.12

Nitrosomonas sp. HPC101 Nitrosomonas sp. DYS323 Nitrosomonas sp. DYS317 Nitrosomonas sp. WH-2 Nitrosomonas europaea Nm50 Nitrosomonas europaea Nm103 Nitrosomonas sp. ENI-11 Nitrosomonas sp. Koll-21 Nitrosomonas sp. IWT514 Nitrosomonas sp. AN1-5 Nitrosomonas halophila Nm1 Nitrosomonas sp. Nm107 Nitrosomonas sp. Nm104 Nitrosomonas mobilis Nc2 Nitrosomonas mobilis Nm93 Nitrosomonas sp. TNO632 Nitrosomonas communis Nm2 Nitrosomonas sp. Nm33 Nitrosomonas sp. Nm41 Nitrosomonas sp. Nm58 Nitrosomonas nitrosa Nm90 Nitrosomonas sp. Nm148

N. communis lineage (Cluster 8) 94.3

Nitrosomonas sp. B19-i-NH4 Nitrosomonas sp. TA 921-i-NH4

N. marina lineage (Cluster 6b) 95.6

Nitrosomonas sp. Nm63 Nitrosomonas sp. C-113 Nitrosomonas sp. Nm51 Nitrosomonas marina Nm22 Nitrosomonas sp. 122 Nitrosomonas sp. C-45 Nitrosomonas sp. NH4W Nitrosomonas sp. N03W Nitrosomonas sp. URW Nitrosomonas aestuarii Nm36 Nitrosomonas ureae Nm10 Nitrosomonas sp. AL212 Nitrosomonas sp. Nm47 Nitrosomonas oligotropha Nm45 Nitrosomonas sp. Nm86 Nitrosomonas sp. Nm59 Nitrosomonas sp. JL21 Nitrosomonas sp. Nm84 Nitrosomonas cryotolerans Nm55 Nitrosomonas sp. TT140-89A Nitrosomonas sp. TT140-098-2 Nitrosomonas sp. Nm143 Nitrosomonas sp. C-17

N. oligotropha lineage (Cluster 6a) 95.8

Nitrosomonas cryotolerans lineage Nitrosomonas sp. Nm143 lineage (Cluster 9) 97.5 Nitrosospira lineage 96.1

0.10

Fig. 7. 16S rRNA based phylogenetic tree of the nitrosomonads. The tree includes all isolates for which 16S rRNA gene sequences longer than 1000 nucleotides are available. Described species are depicted in bold. Maximum likelihood, maximum parsimony, and neighbor-joining trees were calculated and merged. Multifurcations connect branches for which a relative order cannot be unambiguously determined by applying different treeing methods. Filled and empty dots indicate parsimony bootstrap values (100 resamplings) above 90% and 70%, respectively. For each cluster, the minimum 16S rRNA sequence similarity between two of its members is depicted. Sequences included in the analysis were published by Head et al. (1993), Suwa et al. (1997), Juretschko et al. (1998), Sorokin et al. (1998), Aakra et al. (1999a), Aakra et al. (1999b), Yamagata et al. (1999), Purkhold et al. (2000), and Purkhold et al. (2002). Sequences of strains CNS326, G1, K1, IWT202, TK794, WH-2, Koll-21, DYS317, DYS323, IWT514, TNO632, marine bacteria C-45, NH4W, 122, URW, NO3W, C-113, TT140-098-2, TT14089A, and estuarine bacteria TA 921-i-NH4, B19-i-NH4, C-17 are unpublished but available at GenBank. Scale bar represents 10% estimated sequence divergence.

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria Nitrosospira sp. Nsp1 Nitrosospira sp. Nsp4 Nitrosospira sp. GS833

Fig. 8

Nitrosospira sp. E12 Nitrosospira sp. TYM9 Nitrosospira sp. F3 Nitrosospira sp. NRS527 Nitrosospira sp. Nsp40 Nitrosospira sp. Nsp41 Nitrosospira sp. Np39-19 Nitrosospira sp. Np22-21 Nitrosospira sp. Nv6 Nitrosospira sp. Nv12 Nitrosospira birensis Nsp10 Nitrosospira sp. Nsp62 Nitrosospira sp. L115

“Cluster 3”

Nitrosospira sp. 24C Nitrosospira sp. AF Nitrosospira sp. A16 Nitrosospira sp. A4 Nitrosovibrio (Nitrosospira) tenuis Nv1 Nitrosolobus (Nitrosospira) sp. NL5 Nitrosospira sp. NpAV Nitrosospira sp. C-141 Nitrosospira sp. Nsp17 Nitrosospira sp. Nsp2 Nitrosospira sp. RY3C Nitrosospira sp. RY6A Nitrosospira sp. TCH716 Nitrosospira sp. PJA1 Nitrosolobus (Nitrosospira) multiformis NL13 Nitrosospira sp. Nsp58 Nitrosospira sp. Nsp57 Nitrosospira sp. O13 Nitrosospira sp. AHB1 Nitrosospira III7 Nitrosospira sp. T7

“Cluster 2”

Nitrosospira sp. B6 Nitrosospira sp. O4 Nitrosospira sp. Ka3 Nitrosospira sp. Ka4 Nitrosomonas

“Cluster 4”

Nitrosospira sp. III2 Nitrosospira sp. D11 Nitrosospira sp. GM4

“Cluster 0”

Nitrosospira sp. 40KI Nitrosospira sp. Nsp12 Nitrosospira sp. Nsp5 Nitrosospira sp. Nsp65

0.10

Fig. 8. 16S rRNA-based phylogenetic tree of the highly related genera Nitrosospira, Nitrosolobus, and Nitrosovibrio. The tree includes all isolates for which 16S rRNA gene sequences longer than 1000 nucleotides are available. Described species are depicted in bold. Maximum likelihood, maximum parsimony, and neighbor-joining trees were calculated and merged. Multifurcations connect branches for which a relative order cannot be unambiguously determined by applying different treeing methods. Filled and empty dots indicate parsimony bootstrap values (100 resamplings) above 90% and 70%, respectively. Sequences included in the analysis were published by Head et al. (1993), Teske et al. (1994), Utaaker et al. (1995), Tokuyama et al. (1997), Aakra et al. (1999a), Aakra et al. (1999b), Aakra et al. (2001b), and Purkhold et al. (2002). Sequences of strains GS833, E12, NRS527, NpAV, RY6A, RY3C, TCH716, and PJA1 are unpublished but available at GenBank. Scale bar represents 10% estimated sequence divergence.

789

see the section on Environmental Sequences in this Chapter). Purkhold et al. (2000) subsequently extended this system by the proposing cluster 0. These nitrosospira clusters are found with all treeing methods but not all of them are well supported by bootstrap analysis. Therefore, clusters of the Nitrosospira lineage are not comparable with the distinct lineages within the Nitrosomonas group. This reflects the fact that the minimum 16S rRNA sequence similarity of 96.1% within the entire Nitrosospira group is higher than those within the individual, described Nitrosomonas lineages (Fig. 7). It should also be noted that DNA-DNA hybridization analyses suggest that the Nitrosospira clusters 0, 2 and 4 might each represent only one single species (H.-P. Koops, unpublished observation). Taken together, it is thus questionable whether subdivision of the Nitrosospira lineage into the above-mentioned clusters or other subdivisions (Aakra et al., 2001b) is justified. Owing to the limited discriminatory power of comparative 16S rRNA analysis, in particular for the Nitrosospira lineage, other phylogenetic marker molecules were tested for phylogeny inference of AOB. Aakra and co-workers determined the 16S–23S rRNA gene intergenic spacer region (ISR) of several AOB and used these sequences for phylogenetic analysis (Aakra et al., 2001b). In that study, the authors postulated highly consistent ISR- and 16S rRNA-based tree topologies for AOB and thus suggested the usefulness of the ISR for future studies on AOB diversity and evolution. However, careful inspection of the phylogenetic trees presented in that publication reveal significant topological incongruencies between 16S rRNA and ISR trees. Owing to the high variability of the ISR, these sequences are very difficult to align and not suitable to reliably determine evolutionary relationships above the species level. Recently, the amoA gene coding for the active site polypeptide of the ammonia monooxygenase has been used as an additional phylogenetic marker molecule for AOB (McTavish et al., 1993; Klotz and Norton, 1995; Rotthauwe et al., 1995; Suwa et al., 1997; Hommes et al., 1998; Alzerreca et al., 1999; Yamagata et al., 1999; Horz et al., 2000; Purkhold et al., 2000; Aakra et al., 2001a; Casciotti and Ward, 2001; Purkhold et al., 2003). PCR primers that allow amplification of a 453bp fragment of this gene are generally used in these studies (Rotthauwe et al., 1997; modified by Stephen et al., 1999). Phylogeny inference based on the deduced amino acid sequence of the amoA gene fragment is overall consistent with the 16S rRNA phylogeny of AOB. Members of the genera Nitrosospira, Nitrosolobus and Nitrosovibrio form a tight monophyletic grouping with no obvious substructure. Within the

790

Fig. 9

H.-P. Koops et al.

CHAPTER 3.2.12

Nitrosomonas eutropha Nm57 Nitrosomonas sp. TK794 Nitrosomonas sp. F6, F5, GH22 Nitrosomonas sp. ENI-11 Nitrosomonas europaea Nm50

N. europaea/ Nc. mobilis lineage (Cluster 7)

Nitrosomonas halophila Nm1 Nitrosomonas sp. AN1 Nitrosomonas mobilis Nc2 Nitrosomonas sp. Nm41 Nitrosomonas sp. Nm58

N. communis lineage (Cluster 8)

Nitrosomonas communis Nm2 Nitrosomonas sp. Nm33 Nitrosomonas nitrosa Nm90 Nitrosomonas sp. Nm148

Nitrosomonas sp. Nm51 Nitrosomonas marina Nm22

N. marina lineage (Cluster 6b)

Nitrosomonas aestuarii Nm36 Nitrosomonas ureae Nm10 Nitrosomonas sp. AL212 Nitrosomonas sp. JL21 Nitrosomonas sp. Nm59 Nitrosomonas sp. Nm84

N. oligotropha lineage (Cluster 6a)

Nitrosomonas sp. Nm86 Nitrosomonas oligotropha Nm45 Nitrosomonas sp. Nm47 Nitrosomonas sp. Nm143

Nitrosomonas sp. Nm143 lineage

Nitrosomonas cryotolerans Nm55

N. cryotolerans lineage

Nitrosospira lineage

nitrosomonads the N. europaea/Nc. mobilis lineage and the N. marina lineage are also found with all treeing methods, while the N. communis and the N. oligotropha lineage are not always monophyletic (Fig. 9). If amoA nucleic acid sequences are used for phylogenetic analysis, basically the same picture emerges with the exception that the N. europaea/eutropha lineage is no longer monophyletic (data not shown). If compared to the 16S rRNA-based phylogeny of AOB, AmoA analysis does provide less resolution, reflecting that a relatively short (151 positions) and highly conserved (93 positions have an identical amino acid in at least 98% of the betaproteobacterial AOB) amino acid sequence stretch is used as marker. The information content of AmoA sequences could be significantly extended in future studies by the development of primers that allow the amplification of more complete amoA gene fragments. First attempts

Fig. 9. AmoA-based phylogenetic tree of the betaproteobacterial AOB. Described species are depicted in bold. The 453-bp gene fragment obtainable with the most commonly used amoA PCR primers (Rotthauwe et al., 1995) was used for phylogeny inference. AmoA sequences shorter than 414 nucleotides were excluded from the analysis. Protein maximum likelihood, protein maximum parsimony, neighbor-joining, and Fitch trees were calculated and merged. Multifurcations connect branches for which a relative order cannot be unambiguously determined by applying different treeing methods. Filled and empty dots indicate parsimony bootstrap values (100 resamplings) above 90% and 70%, respectively. Sequences included in the analysis were published by McTavish et al. (1993), Holmes et al. (1995), Rotthauwe et al. (1995), Suwa et al. (1997), Yamagata et al. (1999), Aakra et al. (2001a), Casciotti et al. (2001), Sorokin et al. (2001), Norton et al. (2002), and Purkhold et al. (2003). Sequences of Nitrosospira sp. C-57 and Nitrosomonas sp. TK794 are unpublished but available at GenBank. Scale bar represents 10% estimated sequence divergence.

0.10

in this direction were recently published by Norton et al. (2002). Correlation plots of amoA and AmoA similarity versus 16S rRNA similarity of all possible pairs of betaproteobacterial AOB isolates (Fig. 10) demonstrate that 1) 16S rRNA is more conserved than amoA or AmoA and 2) AOB showing below 80% amoA nucleic acid similarity (or 85% AmoA amino acid similarity) always possess less than 97.0% 16S rRNA sequence similarity (the currently accepted 16S rRNA threshold value for bacteria genospecies assignment; Stackebrandt and Goebel, 1994). AmoA sequences of a new AOB isolate with less than 80% nucleic acid similarity (or 85% amino acid similarity) to described AOB species are thus indicative of a previously undiscovered species. An amoA or AmoA sequence with a higher similarity to a described AOB species can represent multiple gene copies, different strains

Fig. 10. Correlation plots of 16S rRNA similarity with amoA (panel A) and AmoA (panel B) similarity values. Solid lines indicate 16S rRNA threshold values for species delineation. Dotted lines indicate the amoA and AmoA threshold values below which amoA and AmoA sequences are indicative of novel AOB species.

The Lithoautotrophic Ammonia-Oxidizing Bacteria

% 16S rRNA gene similarity

CHAPTER 3.2.12

100 99 98 97 96 95 94 93 92 91 90 89 88 87 86

791

multiple copies of the same strain different strains of the same species different species

55

60

65

70

75

80

85

90

95 100

% 16S rRNA gene similarity

% amoA similarity

100 99 98 97 96 95 94 93 92 91 90 89 88 87 86

multiple copies of the same strain different strains of the same species different species

55

60

65

70

75

80

85

90

95 100

% AmoA similarity

of this species, or a novel AOB species. The latter possibility exists since 16S rRNA similarities between different species can be higher than 97%.

Environmental Sequences Molecular diversity surveys of AOB almost exclusively focussed on betaproteobacterial AOB. Stephen et al. (1996) were the first to investigate the diversity of these bacteria in an environmental sample using directly retrieved 16S rRNA gene sequences (see the section on Molecular Techniques in this Chapter). Since then, AOB species richness of numerous natural and engineered systems has been analyzed by the 16S rRNA approach. However, the PCR primers applied in several of these studies do not cover all recognized AOB and thus probably provided a biased view of the natural AOB diversity and their environmental distribution (see the

section on Molecular Techniques in this Chapter). Interestingly, of the 508 16S-rRNA clones affiliated with betaproteobacterial AOB (status August 2002), more than 83% cluster with cultured representatives of this guild. The remaining clones form two novel clusters, one within the nitrosomonads (environmental lineage 5) and one within the nitrosospiras (cluster 1; Fig. 11). On the basis of the results of the various AOB 16S rRNA diversity surveys, environmental distribution patterns of the different AOB clusters can be revealed and compared to the findings obtained with cultivation-based methods (Fig. 12). A more detailed discussion of environmental distribution patterns of the different AOB is provided in a separate section (see the section on Distribution of AOB in Nature in this Chapter). The natural diversity of ammonia oxidizers in various environments also has extensively been studied by comparative sequence analysis

792

H.-P. Koops et al.

CHAPTER 3.2.12

Isolates Enrichments 24 59

Nitrosospira

43

“Cluster 0”

6

-

18

“Cluster 1”

-

-

59

“Cluster 2”

6

2

35

“Cluster 3”

31

23

118

“Cluster 4”

3

1

30

2

-

-

1 -

8

16

8

12

92

2 4 6

6 1 2

4 1 2

5

1

65

14

3

50

172

34 2 14 112

Nitrosospira sp. Nsp57 + Nsp58 Nitrosospira sp. Nsp65 environmental clones N. oligotropha, N. ureae + RS

10

Nitrosospira sp. + RS N. marina + RS N. aestuarii + RS

71

N. europaea + RS

12 6

Nitrosomonas

Environ clones

67

N. eutropha + RS

env. lineage 5 N. oligotropha lineage

N. marina lineage

N. europaeal Nc. mobilis lineage

6

N. halophila + RS

6

-

2

12

N. mobilis + RS

5

2

5

7

N. communis + RS

6

-

2

21

Nitrosomonas sp. Nm143 + RS

4

8

9

1

-

-

110

69

508

Nitrosomonas cryotolerans

Â

Fig. 11. Schematic 16S rRNA-based phylogenetic classification of the betaproteobacterial AOB. Multifurcations connect branches for which a relative order could not be unambiguously determined by applying different treeing methods. The height of each triangle represents the number of sequences in the cluster. It should be noted that 16S rRNA sequences shorter than 1000 nucleotides were only considered if they could be unambiguously assigned to one of the lineages. All other short 16S rRNA sequences had to be omitted because their phylogenetic analysis cannot reliably be performed (Ludwig et al., 1998). The 16S rRNA sequences used in this analysis were published by Takahashi et al. (1992), Head et al. (1993), McCaig et al. (1994), McCaig et al. (1999), Teske et al. (1994), Rotthauwe et al. (1995), Utaaker et al. (1995), Pedersen et al. (1996), Stephen et al. (1996), Kowalchuk et al. (1997), Kowalchuk et al. (1998), Kowalchuk et al. (2000a), Kowalchuk et al. (2000b), Suwa et al. (1997), Tokuyama et al. (1997), Juretschko et al. (1998), Logemann et al. (1998), Princic et al. (1998), Speksnijder et al. (1998), Aakra et al. (1999a), Aakra et al. (1999b), Aakra et al. (2000), Aakra et al. (2001b), Bruns et al. (1999), Mendum et al. (1999), Philips et al. (1999), Philips et al. (2000), Radeva et al. (1999), Whitby et al. (1999), Whitby et al. (2001a), Whitby et al. (2001b), Yamagata et al. (1999), Abd El Haleem et al. (2000), Bano et al. (2000), Ward et al. (2000), Bollmann et al. (2001), Burrell et al. (2001), Chang et al. (2001), de Bie et al. (2001), Daims et al. (2001a), Gieseke et al. (2001), Smith et al. (2001), Sorokin et al. (2001), Hollibaugh et al. (2002), Nicolaisen et al. (2002), Regan et al. (2002), and Purkhold et al. (2003). In addition, unpublished 16S rRNA sequences (Accession numbers: AF107527, AJ441258, AJ441259, AJ441260, AJ441261. AJ441262, AJ441263, AJ441264, AJ441265, AJ441266] AJ441267, AJ441268, AJ441269, AJ441270, AJ441271, AJ441272], AJ441273, AJ441274, AJ441275, AJ441276, AJ441277, AJ441278, AJ441279, AJ441280, AJ441281, AJ441282, AJ441283, AJ441284, AJ441285, AF034139, AF034140, AF034141], AF034142, AF034143, AF034144, AF034147, AJ318197, U57617, AF510862, AF510863, AF510864, AF510865, AJ245751, AJ245752, AJ245753, AJ245754, AJ245755, AJ245756, AJ245757, AJ245758, AJ245759, AJ245760, AJ431350, AJ431351], AF414581, Y10128, Y10127, AJ224941, AF353155, AF353156, AF353157, AF353158, AF353159, AF353161, AF353162, AF353163, AF353164, AF359341, AF363287, AF363289, AF363290, AF363291, AF363292, AF363293, and A036898) available in GenBank were analyzed. RS: related sequences.

of environmentally retrieved amoA clones (Rotthauwe et al., 1997; Helmer et al., 1999; Stephen et al., 1999; Baribeau et al., 2000; Horz et al., 2000; Ivanova et al., 2000; Nold et al., 2000; Purkhold et al., 2000; Schmid et al., 2000; Chang et al., 2001; Gieseke et al., 2001; Oved et

al., 2001; Dionisi et al., 2002; Sakano et al., 2002). In total, 383 amoA clones are currently available from these studies (status August 2002). Consistent with the 16S rRNA-based AOB diversity surveys, most amoA clones obtained are affiliated with sequence clusters

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

Nitrosospira “Cluster 0”

Clones/Enr. (4 habitats)

Nitrosospira “Cluster 1”

Clones/Enr. (8 habitats)

Nitrosospira “Cluster 2”

Clones/Enr. (10 habitats)

Nitrosospira “Cluster 3”

Clones/Enr. (26 habitats)

Nitrosospira “Cluster 4”

Clones/Enr. (10 habitats)

enironmental lineage 5

Clones/Enr. (7 habitats)

N. oligtropha lineage

Clones/Enr. (15 habitats)

N. marina lineage

Clones/Enr. (9 habitats)

N. europaeal Nc. mobilis lineage

Clones/Enr. (18 habitats)

N. communis lineage Nitrosomonas sp. Nm143 lineage

793

Isolates (6 habitats)

Isolates (0)

Isolates (3 habitats)

Isolates (20 habitats)

Isolates (3 habitats)

Isolates (0)

Isolates (9 habitats)

Isolates (10 habitats)

Isolates (12 habitats)

Isolates (6 habitats) Clones/Enr. (1 habitats) Isolates (4 habitats) Clones/Enr. (6 habitats)

0%

10%

20%

30%

40%

50%

60%

70%

80%

soil (n=34, m=19)

marine environment (n=13, m=8)

wastewater treatment plant (n=15, m=6)

coastal sand dune (n=0, m=2)

freshwater environment (n=4, m=10)

estuary (n=3, m=2)

90%

100%

others (n=4, m=3)

Fig. 12. Environmental distribution patterns of AOB clusters. For each cluster, the origin (sampling site) of each 16S rRNA sequence belonging to this cluster was determined. Subsequently, we counted the number of different sampling sites in which members of a particular cluster were detected. This number of habitats, which is given in parentheses, was set to 100%. In the next step, the sampling sites were grouped into the categories wastewater treatment plants, soil (including a rhizoremediation plant), freshwater (including sediment), marine (including sediment), estuary, and coastal sand dune. All other habitats were lumped together. For each cluster the proportion of each category is shown. For comparison, an identical analysis was performed of the available isolates for each cluster. The displayed legend explains the color-coding of the habitats. In addition, the number of different samples of a particular habitat category analyzed by the 16S rRNA approach (n) or cultivation (m) is given. For example, 16S rRNA AOB clones are available from 34 different soil samples and 19 soils were examined by AOB cultivation.

defined by cultured ammonia oxidizers (Table 4). The environmental distribution patterns of AOB inferred from AmoA analysis (Fig. 13) confirm the results from the 16S rRNA studies (Fig. 12), although the limited resolution provided by AmoA does not allow specific data for the individual Nitrosospira clusters to be extracted or the N. oligotropha to be distinguished from the N. marina cluster. According to the threshold values inferred from the amoA/ 16S rRNA correlation plots (Fig. 10), only two (clone GLII-9 from a wastewater treatment plant [wwtp] and clone Pluß see from a lake) of the environmental amoA clones probably represent novel AOB species. These observations confirm that ammonia oxidizers already deposited in culture collections might be surprisingly representative of the natural diversity within this guild. However, it is important to stress that the sequences from 16S rRNA or amoA gene libraries from environmental samples rarely

contain sequences completely identical to those of cultured organisms. Since no DNA-DNA hybridization data can be obtained for noncultured organisms, it can, according to the current species definition in bacteriology (Stackebrandt et al., 1988), not be decided whether these sequences represent novel or known ammonia oxidizer species. In this context, one should however bear in mind that PCR and cloning procedures introduce microvariation artifacts in cloned sequences (Speksnijder et al., 2001) and thus might cause (at least a part of) the high degree of microheterogeneity observed for AOB sequence clusters detected in environmental 16S rRNA and amoA clone libraries. Although the 16S rRNA and amoA library approaches provided valuable insights into the natural diversity of AOB, the dependency of these techniques on nucleic acid extraction, PCR and cloning heavily biases the results (Wintzigerode and Goebel, 1997). Understand-

794

H.-P. Koops et al.

CHAPTER 3.2.12

Table 4. Phylogenetic affiliation of amoA sequences obtained from environmental samples. AmoA-sequences analyzed for this Table were obtained from Baribeau et al. (2000), Chang et al. (2001), Dionisi et al. (2002), Gieseke et al. (2001), Ivanova et al. (2000), Helmer et al. (1999), Henckel et al. (1999), Holmes et al. (1999), Horz et al. (2000), Nold et al. (2000), Oved et al. (2001), Purkhold et al. (2000), Purkhold et al. (2003), Rotthauwe et al. (1997), Sakano et al. (2002), Schmid et al. (2000), and Stephen et al. (1999). In addition, unpublished environmental amoA sequences determined by the authors or available in GenBank (Accession numbers: AJ277459], AF338319, AF338320], AF239878, AF239879, AF239880, AF239881, AF239882, AF239883, AF239884], AJ388568, AJ388569, AJ388570, AJ388571, AJ388572, AJ388573, AJ388574, AJ388575, AJ388576, AJ388577, AJ388578, AJ388579, AJ388580, AJ388581, AJ388582, AJ388583, AJ388588, and AJ388589) were used.

Nitrosospira cluster N. marina and N. oligotropha cluster N. europaea/Nc. mobilis cluster N. communis cluster Nitrosomonas sp. Nm143 cluster N. cryotolerans cluster Σ

Total

Isolates

Enrichments

Environmental clones

152 116 156 37 1 1 463

37 16 16 6 1 1 77

3 0 0 0 0 0 3

112 100 140 31 0 0 383

Nitrosospira lineage (23) N. oligotropha + N. marina lineage (24)

N. communis lineage (9)

N. europaeal Nc.mobilis lineage (23)

0%

20%

40%

60%

80%

soil (including rice roots, n=13)

marine environment (n=2)

wastewater treatment plant (n=22)

estuary (n=2)

freshwater environment (n=7)

biofilter (n=1)

100%

Fig. 13. Environmental distribution patterns of AOB clusters based on AmoA analysis. For each cluster, the origin (sampling site) of each AmoA sequence belonging to this cluster was determined. This analysis only included sequences from enrichments and sequences directly retrieved from the environment. Subsequently, we counted the number of different sampling sites in which members of a particular cluster were detected. This number, which is given in parentheses after the cluster designation, was set to 100%. In the next step, the sampling sites were grouped into the categories wastewater treatment plants, soil, freshwater (including sediment), estuary, biofilter, and marine (including sediment). For each cluster, the proportion of each category is shown. The displayed legend explains the color-coding of the habitats. In addition, the number of different sampling sites of a particular habitat category analyzed by the 16S rRNA approach (n) is given. Please note that the depicted grouping does not reflect the phylogenetic relation but results from the poor resolution of the amplified amoA gene fragment.

ing the ecology of AOB requires quantitative data on their in situ abundance and activity in ecosystems. The AOB community composition in a system can be quantitatively analyzed by using rRNA-targeted probes in combination with dot blot or in situ hybridization techniques (see the section on Molecular Techniques in this Chapter). However up to now, the AOB composition of only wastewater treatment reactors and full-scale plants has been characterized with these methods (Wagner et al., 1995; Mobarry et al., 1996; Juretschko et al., 1998;

Schramm et al., 1999; Schramm et al., 2000; Daims et al., 2001b).

Methods Useful for In Situ Detection of AOB Classical Techniques Traditionally, the abundance, diversity and ecology of AOB were investigated by cultivationdependent methods. For the enumeration of

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

viable AOB, the most probable number (MPN) technique often has been used, sometimes in parallel with other techniques. The efficiency of MPN has been calculated from comparisons with results obtained with fluorescent antibody (FA) counting (Belser, 1979; Ward, 1982) or calculations of activity per cell (Belser and Mays, 1982). Results obtained from these comparisons have led to the suggestion that the MPN efficiency may range between 0.001 and 0.05% (Hall, 1986). One important reason of underestimating AOB with MPN may be that AOB often form cell aggregates in the environment (Wagner et al., 1995; Stehr et al., 1995b; Juretschko et al., 1998; Koops and Pommerening-Röser, 2001). Another reason may be that the media used in MPN counting of AOB often have not been well adapted (ammonia concentration, salinity, and pH value) to the conditions in the environment under investigation (Koops and PommereningRöser, 2001). Qualitative characterization of natural AOB populations is a very time consuming process. This especially holds true if classical methods are applied. The isolation of numerically dominant species will take at least some months (see the section on Enrichment, Isolation, and Maintenance in this Chapter). Moreover, considerable experience in working with AOB in the laboratory is required. However, this approach is currently the only option that allows the physiological properties of those species representing the numerically dominant AOB population in the respective environment to be completely investigated. These findings can help determine why distinct species of AOB are dominant in certain environments. Phenotypic identification of new isolates of AOB is possible if the strain belongs to one of those species already being described in the literature. Useful parameters are the physiological properties (Ks values of ammonia-oxidation, requirement or tolerance of increased salt concentrations, and possession of urease), morphological characteristics (shape and size of cells, arrangement of intracytoplasmic membranes, and possession of carboxysomes), and G+C content of the DNA. For unambiguous identification, DNA-DNA hybridization with the type strain of the respective species should be carried out.

Immunological Techniques In the 1970s, the cultivation-dependent approach was complemented by the fluorescent antibody technique, which allowed (for the first time) members of this guild to be detected in situ. Generally, polyclonal antibodies against several AOB

795

were produced by injecting rabbits with the respective cells. Several studies demonstrated in situ visualization of AOB in environmental samples using these antisera for immunofluorescence and claimed that this approach allows a more rapid and accurate enumeration of AOB compared to enrichment culture methods (Belser, 1979; Ward and Perry, 1980; Ward and Carlucci, 1985; Abeliovich, 1987; Völsch et al., 1990). Although the fluorescent antibody (FA) approach represented a significant step forward in the history of research on nitrifier ecology, this technique has several disadvantages. Most importantly, pure cultures of AOB from the environment under investigation are required to produce the antibodies, since different serological groups exist within at least some of the AOB species. Therefore, the FA-technique, although allowing direct in situ detection of AOB, is indirectly dependent on successful cultivation of the target organisms. Consequently, this technique will not allow identification of novel, previously uncultured AOB species. Furthermore, the various polyclonal antibodies raised in different laboratories against different AOB are not easily available and thus their use is often restricted to a few laboratories. Another major disadvantage of the FA-technique is that the specificity of the obtained antibodies cannot be tailored in silico (i.e., using a computer) and must be determined in time-consuming experimental screening assays using different pure cultures. Finally, the FA-technique potentially suffers from false negative and false positive signals. The false negative signals might result from penetration problems of the relatively large antibodies in dense flocs or biofilms. False-positive signals were described as a result from nonspecific antibody binding to extracellular material or detritus (Szwerinski et al., 1985). Recently, polyclonal antibodies against the purified β subunit of the ammonia monooxygenase (AmoB) of Nitrosomonas eutropha were shown to target all betaproteobacterial AOB by Western blot analysis (Pinck et al., 2001). After appropriate AOB cell permeabilization (Bartosch et al., 1999), the α AmoB antibodies are also suitable for immunofluorescence-based group-specific detection of betaproteobacterial AOB (E. Spieck, personal communication). Since these polyclonal antibodies target a key enzyme of the betaproteobacterial AOB, this assay is expected to be highly specific and will probably also allow yet unknown members of this group to be targeted. In the future, this technique most probably can be standardized to become the first choice method for general counting of betaproteobacterial AOB in those environmental samples that are well suited for FA application.

796

H.-P. Koops et al.

Molecular Techniques After the establishment of 16S rRNA-based phylogeny of cultured AOB (Head et al., 1993; Teske et al., 1994; Pommerening-Röser et al., 1996; see the section on Phylogeny of AOB in this Chapter), many different 16S rRNA-targeted PCR primers and probes for dot blot or fluorescence in situ hybridization (FISH) have been developed (Table 5). Most frequently the natural diversity of AOB has been investigated by comparative analyses of PCR amplified and cloned AOB 16S rRNA gene fragments (see the section on Distribution of AOB in Nature in this Chapter). Apparently, the specificity of the applied PCR primers dramatically determines the outcome of such molecular AOB diversity surveys. In this context, it is important to note that the actual specificity of many primers used in previous investigations significantly differs from the indicated (intended) specificity listed in the original publications (Tables 6 and 7). These inconsistencies were detected after re-evaluating the primer/probe specificities based on the current, encompassing AOB 16S rRNA gene database. For example the PCR primer NitA, which is frequently used for AOB diversity surveys (Abd El Haleem et al., 2000; Ward et al., 2000; Hollibaugh et al., 2002), shows mismatches to all cultured nitrosospiras, all members of the N. communis and N. oligotropha lineage, and most members of the N. marina lineage. To avoid such pronounced biases, the use of PCR primers with high sensitivity (targeting all known or at least most betaproteobacterial AOB) but relatively low specificity (not perfectly excluding other bacteria) is recommended. Furthermore, the amplified 16S rRNA gene fragment must have a size larger than 1000 nucleotides to allow reliable phylogeny inference. The above-mentioned criteria are currently best fulfilled by the β AMOf and β AMOr primer pair. This approach accepts the amplification of non-AOB 16S rRNA gene fragments, which subsequently have to be identified by phylogenetic analysis or hybridization with probes with excellent specificity (e.g., probe Nso1225). Recently, the toolbox for AOB detection has been extended by analyzing PCR amplified 16S rRNA gene fragments with denaturing gradient gel electrophoresis (DGGE; Kowalchuk et al., 1997; Nicolaisen and Ramsing, 2002). This fingerprinting approach allows a rapid comparative assessment of AOB diversity in many samples but requires subsequent controls like cloning and sequencing of bands or hybridization of membrane-blotted DGGE gels with suitable probes. Furthermore, the length of PCR amplificates that can be analyzed is limited to approximately 500 bp, which limits primer selection and hampers phylogeny inference.

CHAPTER 3.2.12

Quantitative analysis of AOB community composition and abundance in the environment requires the use of methods not affected by PCR biases (Wintzigerode and Goebel, 1997). For this purpose, both dot blot and FISH methods have been developed and applied for AOB diversity research (Wagner et al., 1995; Mobarry et al., 1996; Schramm et al., 1996; Juretschko et al., 1998; Okabe et al., 1999; Liebig et al., 2001; Nogueira et al., 2002). The major advantage of dot blot analysis is that it can be applied to almost all environmental samples, while FISH does not work quantitatively in samples having a high autofluorescence or containing nontransparent particles (like soils). On the other hand, FISH allows visualization of the target bacteria and quantification of the AOB community composition and absolute AOB cell numbers (Daims et al., 2001c), while dot blot analysis only provides information on the relative or absolute abundance of probe target rRNA molecules. Whether, for example, a high rRNA content of a certain AOB species in an environmental sample originates from many target cells with a low rRNA concentration or a few cells with a high rRNA concentration cannot be figured out by dot blot hybridization. It has been postulated that the rRNA content of a bacterial cell is directly related to its physiological activity. It must however be noted that this link does not always hold true for AOB. Quantitative FISH experiments showed that AOB have high cellular ribosome contents even after prolonged complete physiological inhibition (Wagner et al., 1995) or starvation (Morgenroth et al., 2000). Therefore, physiological activity of AOB cannot reliably be inferred from the above-mentioned methods and must be determined for example by combining FISH and microautoradiography (Lee et al., 1999), FISH and microelectrodes (Schramm et al., 1999), or stable isotope probing (Whitby et al., 2001a). In addition to rRNA-based methods, the gene encoding the active site polypeptide of the ammonia monooxygenase (amoA) is frequently used as phylogenetic marker for AOB in environmental diversity surveys. Like the 16S rRNA database, the database of amoA sequences of cultured AOB is encompassing (see the section on Phylogeny of AOB in this Chapter). Several primer pairs for specific amplification of amoA have been published (Holmes et al., 1995; Rotthauwe et al., 1995; Sinigalliano et al., 1995; Mendum et al., 1999; Webster et al., 2002) but only the primer pair described by Rotthauwe et al. (1995), amplifying 55% of the amoA gene and slightly modified later by Stephen et al. (1999), has been shown to be suitable for detecting all tested betaproteobacterial AOB (Purkhold et al., 2000). The amoA approach for

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

797

Table 5. List of published 16S rRNA-targeted primers and probes. Probes that have been successfully used for FISH are shaded. Listed are specificities of primers and probes as given in the original publications. For an overview of the actual number of mismatches of each primer or probe with the 16S rRNA of all ammonia-oxidizer isolates please refer to Tables 6 and 7. Probe/primer

Target site

Specificitya

NM-75 NS-85 NmIIb NSMR32f NSMR71f NSMR34 NSMR76b NitA βAMOf NSPM Nm0 Nsm 156b NmVb CTO189f A/B-GC CTO189f C-GC Nso 190b Noli191b TAOfwd NM198 NmoCL6a_205 NmI Nmo218b TMP1 β-AO233 NspCL1_249 Nmo254a Nmo254 RT1r AAO258 Primer 356f NmoCL6b_376 Nsp436 NmoCL7_439 Nm439 NitD NMOB1f NSMR52f Nsv 443b NspCL4_446 Nsp0 NspCL3_454 NspCL2_458 Nlm 459r NSM1B Primer 517r TAOrev CTO654r NITROSO4Eb NEUb Amβ NitF NitC NmIII NSMR53r NSMR74r NMOB1r NSMR33r RNM-1007 NS-1009 NmIVb NitB Nso 1225b βAMOr Nse 1472b

67–86 76–95 120–139 125–141 126–143 131–149 132–149 136–158 142–162 145–162 148–165 155–173 174–191 189–207 189–207 189–207 191–208 192–208 198–218 205–221 210–225 218–236 226–253 233–249 249–266 254–271 254–271 283–304 258–277 356–372 376–392 436–453 439–456 439–459 439–461 442–461 443–461 443–461 446–463 452–469 454–471 458–475 458–477 478–494 517–533 632–649 632–653 638–657 651–668 738–758 844–862 846–862 998–1018 999–1017 1000–1017 1006–1026 1006–1021 1005–1028 1007–1026 1004–1022 1213–1233 1224–1243 1295–1314 1472–1489

Terrestrial Nitrosomonas spp./Nitrosococcus mobilis Nitrosospira spp. Nitrosomonas communis lineage Nitrosospira tenuis-like AOB Nitrosomonas marina-like AOB Nitrosospira tenuis-like AOB Nitrosomonas marina-like AOB βAOB βAOB βAOB Nitrosomonas spp. Nitrosomonas spp./Nitrosococcus mobilis Nitrosococcus mobilis βAOB βAOB βAOB Nitrosomonas cluster 6a Terrestrial AOB Nitrosomonas ureae + Nitrosomonas sp. AL212 Nitrosomonas cluster 6a Nitrosomonas europaea-lineage Nitrosomonas oligotropha-lineage AOB βAOB Nitrosospira cluster 1 All Nitrosomonas All Nitrosomonas AOB Terrestrial βAOB Nested PCR in NitAB amplicons Nitrosomonas cluster 6b All Nitrosospira Nitrosomonas cluster 7 Nitrosomonas ureae + Nitrosomonas sp. AL212 Nitrosomonas europaea Nitrosococcus mobilis-like AOB Nitrosomonas europaea-like AOB Nitrosospira spp. Nitrosospira cluster 4 Nitrosospira spp. Nitrosospira cluster 3 Nitrosospira cluster 2 Nitrosospira multiformis/Nitrosospira sp. C-141 Nitrosomonas europaea-lineage/Nitrosococcus mobilis Nested PCR in NitAB amplicons Terrestrial ammonia oxidizers βAOB βAOB Most halophilic and halotolerant Nitrosomonas βAOB βAOB βAOB Nitrosomonas marina-lineage Nitrosomonas europaea-like AOB Nitrosomonas marina-like AOB Nitrosococcus mobilis-like AOB Nitrosospira tenuis-like AOB Terrestrial Nitrosomonas spp. Nitrosospira spp. Nitrosomonas cryotolerans-lineage βAOB βAOB βAOB Nitrosomonas europaea-lineage

References Hiorns et al., 1995 Hiorns et al., 1995 Pommerening-Röser et al., 1996 Burrell et al., 2001 Burrell et al., 2001 Burrell et al., 2001 Burrell et al., 2001 Voytek and Ward, 1995 McCaig et al., 1994 Silyn-Roberts and Lewis, 2001 Pommerening-Röser et al., 1996 Mobarry et al., 1996 Pommerening-Röser et al., 1996 Kowalchuk et al., 1997 Kowalchuk et al., 1997 Mobarry et al., 1996 Gieseke et al., 2001; Rath, 1996 Chandler et al., 1997 Suwa et al., 1997 Stephen et al., 1998 Pommerening-Röser et al., 1996 Gieseke et al., 2001 Hermansson and Lindgren, 2001 Stephen et al., 1998 Stephen et al., 1998 Stephen et al., 1998 Stephen et al., 1998 Hermansson and Lindgren, 2001 Hiorns et al., 1995 Hollibaugh et al., 2002 Stephen et al., 1998 Stephen et al., 1998 Stephen et al., 1998 Suwa et al., 1997 Ward et al., 1997 Burrell et al., 2001 Burrell et al., 2001 Mobarry et al., 1996 Stephen et al., 1998 Pommerening-Röser et al., 1996 Stephen et al., 1998 Stephen et al., 1998 Hastings et al., 1997 Hovanec and DeLong, 1996 Hollibaugh et al., 2002 Chandler et al., 1997 Kowalchuk et al., 1997 Hovanec and DeLong, 1996 Wagner et al., 1995 Utaaker and Nes, 1998 Ward et al., 1997 Voytek and Ward, 1995 Pommerening-Röser et al., 1996 Burrell et al., 2001 Burrell et al., 2001 Burrell et al., 2001 Burrell et al., 2001 Hiorns et al., 1995 Hiorns et al., 1995 Pommerening-Röser et al., 1996 Voytek and Ward, 1995 Mobarry et al., 1996 McCaig et al., 1994 Juretschko et al., 1998

a Listed are specificities of primers and probes as given in the original publications. For an overview of the actual number of mismatches of each primer or probe with the 16S rRNA of all ammonia-oxidizer isolates, please refer to Tables 6 and 7. b Probe has been successfully used for FISH.

5 >10 0 0 0 0 3 0 >10 >10 5 2 2 7 3 2 >10 8 9 1 0 >10 >10 >10 >100 >100 >100 >100 >100 >100 4 0 1 0 0 0 2 >10 1 1 4 1 >10 >100 5 3 >10 3 >10 0 1 0 0 1 0 3 0 1 0 >10 4 >100 1

>10 6 0 0 0 0 0 0 7 0 1 2 0 2 0 2 3 0 0 0 0 5 >10 1 >100 >100 >10 >10 >100 >100 0 0 0 0 0 0 1 >10 1 0 0 1 6 >100 2 4 2 0 1 0 0 1 0 0 0 0 0 1 0 5 2 >10 0

NM-75 NS-85 NmII NSMR32f NSMR71f NSMR34 NSMR76 NitA bAMOf NSPM Nm0 Nsm 156 NmV CTO189f A/B-G CTO189f C-GC Nso 190 Noli191 TAOfwd NM198 NmoCL6a_205 NmI Nmo218 TMP1 b-AO233 NspCL1_249 Nmo254a Nmo254 AAO258 RT1r primer 356f NmoCL6b_376 Nsp436 NmoCL7_439 Nm439 NitD NMOB1f NSMR52f Nsv 443 NspCL4_446 Nsp0 NspCL3_454 NspCL2_458 Nlm 459r NSM1B primer 517r TAOrev CTO654r NITROSO4E NEU Amb NitF$ NitC$ NmIII NSMR53r NSMR74r NMOB1r NSMR33r RNM-1007 NS-1009 NmlV* NitB Nso 1225 bAMOr Nse 1472

1MM

0MM

Primer/Probe

NTO NTO

Cluster 2

2

Nsp Nsp AHB III2 40KI D11 GM4 III7 B6 T7 O13 O4 12 5 1

Cluster 0 Nsp 1

Nsp 4

GS 833 F3 E12

TYM 9

NRS 527

Nsp 40

Nsp 10 Nsp 62

1 1

Np 22

Cluster 3

3

6 4

6 4

NL 13

1 1

1

PJ TCH A1 716

1 1

1

1 1 1 1 1

RY 6A

1 1

1

2

1

RY 3C Ka3

Ka4

Cluster 4 Nsp Nsp 57 58

Nsp 65

H.-P. Koops et al.

3

2

2

1

1

1 1 1

Np Nv Nv L C Nsp Nsp Nsp A4 AF A16 24C NL5 Nv1 3919 6 12 115 141 AV 2 17

most sequences in cluster 1 are targeted

Nsp 41

Nitrosospira Isolates

Table 6. Specificity and sensitivity of published 16S rRNA (gene) targeting PCR primers and hybridization probes for nitrosospiras. Abbreviations: NTO, number of non-target organisms outside the AOB with no mismatch (0 MM) or one mismatch (1 MM) in its 16S rRNA (gene) to the respective primer/probe.a

798 CHAPTER 3.2.12

5 >10 0 0 0 0 3 0 >10 >10 5 2 2 7 3 2 >10

8 9 1 0 >10 >10 >10 >100 >100 >100 >100 >100 >100 4 0 1 0 0 0 2 >10 1 1 4 1 >10 >100 5 3 >10 3 >10 0 1 0 0 1 0 3 0 1 0 >10 4 >100 1

>10 6 0 0 0 0 0 0 7 0 1 2 0 2 0 2 3

0 0 0 0 5 >10 1 >100 >100 >10 >10 >100 >100 0 0 0 0 0 0 1 >10 1 0 0 1 6 >100 2 4 2 0 1 0 0 1 0 0 0 0 0 1 0 5 2 >10 0

NM-75 NS-85 NmII NSMR32f NSMR71f NSMR34 NSMR76 NitA bAMOf NSPM Nm0 Nsm 156 NmV CTO189f A/B-GC CTO189f C-GC Nso 190 Noli191 TAOfwd NM198 NmoCL6a_205 NmI Nmo218 TMP1 b-AO233 NspCL1_249 Nmo254a Nmo254 AAO258 RT1r primer 356f NmoCL6b_376 Nsp436 NmoCL7_439 Nm439 NitD NMOB1f NSMR52f Nsv 443 NspCL4_446 Nsp0 NspCL3_454 NspCL2_458 Nlm 459r NSM1B primer 517r TAOrev CTO654r NITROSO4E NEU Amb NitF$ NitC$ NmIII NSMR53r NSMR74r NMOB1r NSMR33r RNM-1007 NS-1009 NmlV* NitB Nso 1225 bAMOr Nse 1472

1MM

0MM

Primer/Probe

NTO NTO

WH 2

Nm ENI- Nm 11 103 50

Koll 21

Nm 57

GH 22

CNS G1 K1 326

IWT 202

IWT 514

1

DYS 317

1

DYS 323

TK F6 F5 794

N. europaea /Nc. mobilis Cluster HPC 101

AN 1-5 Nm 1

Nc 2

Nm Nm 93 104

Nm 107

TNO 632

1

Nm 90

Nm Nm 148 2

Nm 33

Nm 41

Nm 58

N. communis Cluster TA 921

3

B19

Nitrosomonas Isolates Nm 51

Nm 63 Nm 22

2 2 2 1 1

C 113

C 45

1

2

1

mar Nm NH4W URW NO3W 122 36

N. marina Cluster Nm 47

2

Nm 45

Nm 10

1

AL Nm 212 80

JL 21

Nm 59

N. oligotropha Cluster Nm 84

1

TT140 TT140 89A 098-2

Nm 143

Nm143 Cluster C17

Table 7. Specificity and sensitivity of published 16S rRNA (gene) targeting PCR primers and hybridization probes for nitrosomonads. Abbreviations: NTO, number of non-target organisms outside the AOB with no mismatch (0 MM) or one mismatch (1 MM) in its 16S rRNA (gene) to the respective primer/probe.a

Nm 55

CHAPTER 3.2.12 The Lithoautotrophic Ammonia-Oxidizing Bacteria 799

800

H.-P. Koops et al.

AOB diversity research impresses by its specificity and sensitivity but suffers from PCR-biases and relatively low phylogenetic information content (see the section on Phylogeny of AOB in this Chapter). Several fingerprinting techniques like gel retardation, DGGE, and terminal restriction fragment length polymorphism (T-RFLP) have been combined with the amoA approach and allow first insights into the diversity of an amoA PCR amplificate without time-consuming cloning and sequencing analysis (Kowalchuk et al., 1998; Horz et al., 2000; Schmid et al., 2000; Nicolaisen and Ramsing, 2002). Furthermore, an in situ PCR technique is available which allows direct detection of the amoA gene in AOB (Hoshino et al., 2001). Significant method development for AOB diversity research is underway. Several quantitative PCR methods for AOB detection in the environment have been developed (Mendum et al., 1999; Dionisi et al., 2002). Furthermore, absolute AOB numbers in complex samples can also be determined by applying a novel FISH method in combination with confocal laser scanning microscopy and digital image analysis (Daims et al., 2001c). In the future, DNA microarrays composed of several hundred rRNA- or amoA -targeted oligonucleotide probes will almost certainly become available for parallel analyses of AOB community structures. Studies on first generation microarrays carrying a relatively limited number of AOB-targeted probes have already been published (Guschin et al., 1997; Wu et al., 2001).

Distribution of AOB in Nature The distribution of AOB in the environment has traditionally been analyzed by cultivationdependent methods (see the section on Classical Techniques in this Chapter). Recently, these studies were complemented by molecular diversity surveys of various man-made and natural systems (see the section on Molecular Techniques in this Chapter). The combination of both approaches is strongly recommended because each approach is characterized by its specific limitations. MPN-based isolation of AOB target the numerically most important AOB but might overlook not yet cultured AOB species that cannot thrive in the cultivation medium offered (although any clear indication for that is missing in the literature). On the other hand, it is important to interpret PCR-based molecular AOB distribution data with caution because the mere retrieval of a 16S rRNA sequence of an AOB proves neither that this organism is abundant nor that it is physiologically active. This problem is particularly severe for AOB (com-

CHAPTER 3.2.12

pared to many other bacteria) because they can survive very long time periods under unfavorable conditions (Johnstone and Jones, 1988; Wilhelm et al., 1998; Pinck et al., 2001). Therefore, molecular studies aiming at identification of dominant AOB species in an environment should focus on PCR amplificates of the highest positive DNA dilution, an approach analogous to the MPN technique (Feray et al., 1999), or even better include quantitative techniques (see the section on Molecular Techniques in this Chapter). Independent of the method used, the distinction between autochthonous and allochthonous AOB of a particular ecosystem is not trivial, making the natural distribution patterns of the distinct AOB species difficult to define. This problem is particularly severe because many of the environments are strongly influenced by human activity that eventually causes transmission of significant amounts of AOB to foreign systems. For example, AOB typically found in activated sludge also might be detectable in freshwater systems, if effluents of a wastewater treatment plant are disposed close to the sampling site. In this context, the importance of a careful selection of representative sampling sites is obvious. It is also interesting to note that isolations of strains via MPN series in general define more restricted distribution patterns of the distinct species of AOB than obtained from comparable molecular investigations. AOB are present in most aerobic environments including rocks and building stones but can also survive in anaerobic systems (see the section on Dominant Populations of AOB in Different Environments in this Chapter). Since ammonia is the essential energy source for these organisms, their distribution in nature is coupled to geological, biological and anthropogenic sources of reduced nitrogen. Consequently, they have adapted to a broad range of different ammonia concentrations in the diverse environments, reflected by different affinity constants for ammonia. This is one of the most important aspects influencing the distribution patterns of AOB in nature (Suwa et al., 1994; Suwa et al., 1997; Stehr et al., 1995a; Koops and Pommerening-Röser, 2001). On the other side, ammonia is a toxic compound. Therefore, the tolerance of increasing ammonia concentrations (Table 3) is another aspect shaping the distribution patterns of AOB (Bollmann and Laanbroek, 2001). Furthermore, the presence or absence of urease activity was observed to be of ecophysiological relevance for AOB. This property is of special importance in acidic environments, where free ammonia is missing as substrate because it is nearly quantitatively ionized to ammonium. Under such conditions, only those AOB species that can use urea as an alternative

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

ammonia source can build up stable populations (De Boer and Laanbroek, 1989; De Boer et al., 1991; Jiang and Bakken, 1999; Burton and Prosser, 2001). Possession of urease seems also to be an essential property for AOB that successfully colonize oligotrophic soils or aquatic environments. Distribution of AOB in nature also is affected by different salt requirements, salt tolerances and salt sensitivities of the distinct species (Table 3). This is of special importance for their distribution patterns in aquatic systems (such as rivers, lakes, estuaries, marine environments and salt lakes) that significantly differ in salinity (Koops et al., 1990; Koops et al., 1991). The AOB are aerobes. Although their oxygen affinity constants are relatively high (Painter, 1986), AOB possess nitrifying activity not only at oxygen saturation but also at extremely low oxygen concentrations (Goreau et al., 1980). Even ammonia oxidation under anaerobic conditions is being discussed (Schmidt and Bock, 1997; Schmidt and Bock, 1998; Zart et al., 2000). However, specific selection of distinct AOB species by different oxygen concentrations has not yet been reported in the literature. Temperature also may be of importance for distribution patterns of AOB in nature. This has been indicated in some publications (Golovacheva, 1976; Jones et al., 1988; Jiang and Bakken, 1999). Beside environments characterized by constant high or low temperatures, such as hot springs or permafrost soils, environments showing pronounced temperature changes, such as rock surfaces, might harbor interesting AOB. However, these systems have not been studied intensively enough to allow general conclusions on their AOB diversity.

The Betaproteobacterial AOB The betaproteobacterial AOB are divided into two main subgroups, the Nitrosomonas group (containing six distinct lineages) and the Nitrosospira lineage (containing the genera Nitrosospira, Nitrosovibrio and Nitrosolobus). Within the Nitrosomonas group, a relatively strong correlation between the phylogenetically defined lineages (see the section on Phylogeny of AOB in this Chapter) and the distribution patterns of the respective species exists. Within the Nitrosospira lineage, this correlation is not as obvious. The NITROSOMONAS Group The N. europaea/Nc. mobilis Lineage The four cultured species of the N. europaea/Nc. mobilis lineage are moderately halophilic and all show

801

striking halotolerance (Table 3). Most of the cultured strains of the three species N. europaea, N. eutropha and Nitrosococcus mobilis (which is phylogenetically a member of the genus Nitrosomonas) have been isolated from wastewater treatment plants and their molecular signatures are also frequently retrieved from these systems (Figs. 12 and 13). In addition, quantitative FISH results demonstrated that Nc. mobilis and N. europaea are dominant in several wastewater treatment plants (Juretschko et al., 1998; Daims et al., 2001b). Some isolates of this lineage were obtained from strongly fertilized agricultural soils. All these environments are more or less eutrophicated and influenced by anthropogenic activity, frequently rendering the conditions more favorable for these AOB. The successful colonization of these systems requires tolerance of increasing ionic strength and, in particular, of high concentrations of toxic compounds such as ammonia. Consequently, these properties are found in all members of these AOB lineages (Table 3). The natural habitat for members of these lineages has not yet been identified. However, in a recent molecular investigation the presence of N. europaea -like and N. eutropha -like organisms in a salt lake was shown (Ward et al., 2000). This is of interest, since strains belonging to the fourth species of the N. europaea/Nc. mobilis lineage, N. halophila, recently could be isolated from salt lakes in Mongolia (Sorokin et al., 2001). Sporadically, isolates of the N. europaea/Nc. mobilis lineage were obtained from the North Sea (N. halophila and N. mobilis), from the river Elbe estuary (N. europaea) and from freshwater sediments (N. europaea and N. eutropha ; Stehr et al., 1995a; Koops and Pommerening-Röser, 2001). Consistent with these findings, molecular investigations detected members of the N. europaea/Nc. mobilis lineage in fertilized soils or natural fresh water environments (Voytek, 1996; Holmes et al., 1999; Campbell, 2000; Horz et al., 2000; Ivanova et al., 2000; Phillips et al., 2000; Burrell et al., 2001; Oved et al., 2001; Smith et al., 2001; Whitby et al., 2001a; Morris et al., 2002; Radajewski et al., 2002; Webster et al., 2002). The N. communis Lineage The N. communis lineage comprises two entities, the N. communis and the N. nitrosa sublineages, which are currently phylogenetically not clearly distinguishable but have significantly different distribution patterns in nature. The N. communis sublineage contains, in addition to N. communis, three other cultivated species, not yet described by name (Fig. 9). The majority of strains of this sublineage were isolated from approximately pH neutral soils, often agriculturally utilized. Since all three species of this sublineage possess relatively high

802

H.-P. Koops et al.

affinity constants for ammonia and lack urease activity, they probably need relatively high ammonia concentrations in the environment. Most likely this is an important reason that these species seem to be missing in oligotrophic natural soils and in substrate-low natural freshwaters. In contrast to the above subgroup, strains of the N. nitrosa subgroup are commonly distributed in more or less eutrophicated freshwaters. Since the affinity constants for ammonia are similar to those of the species of the N. communis sublineage, the possession of urease might be responsible for this different distribution. Strains of N. nitrosa also have been isolated from marine environments, a fact which is in apparent contrast to the relatively low salt tolerance of these strains observed in laboratory studies (Table 3). Surprisingly, only two 16S rRNA gene clones of the N. communis lineage have been detected in environmental diversity surveys (Gieseke et al., 2001; Fig. 11). This low retrieval rate may have been caused by the fact that many of the commonly applied primers possess mismatches with the 16S rRNA gene of members of this lineage (Table 7). In contrast, amoA sequences affiliated with this lineage were detected in soil, freshwater and wastewater treatment plants (Ivanova et al., 2000; Purkhold et al., 2000; Gieseke et al., 2001; Oved et al., 2001; Dionisi et al., 2002; Nogueira et al., 2002; Fig. 11). The N. oligotropha Lineage The N. oligotropha lineage comprises several species, although only N. oligotropha and N. ureae are described by name. A total of six species of this lineage is available in culture (Fig. 7), and the existence of more species has been indicated by molecular environmental studies (Speksnijder et al., 1998) and several unpublished sequences publicly accessible via GenBank. The majority of cultured strains have been isolated from rivers and lakes (Koops and Pommerening-Röser, 2001). Some isolates originate from sewage disposal plants (Koops and Harms, 1985; Suwa et al., 1997) and from soil samples. The 16S rRNA sequences of this cluster were frequently detected in soil and freshwater (Stephen et al., 1996; Speksnijder et al., 1998; Bruns et al., 1999; Radeva et al., 1999; Kowalchuk et al., 2000b; Phillips et al., 2000; Bollmann and Laanbroek, 2001; Whitby et al., 2001b; Regan et al., 2002). Among the cultivated species of AOB, the representatives of the N. oligotropha lineage reveal the lowest affinity constants for ammonia, being in the range of a few µM. All investigated strains are urease positive. These observations are in accordance with the generally low ammonia concentration in their natural habitats. It is also notable, that representatives of this lineage often occur floc- or biofilm-attached in the environ-

CHAPTER 3.2.12

ment (Stehr et al., 1995a), as well as in laboratory studies (Bollmann and Laanbroek, 2001). Consistent with this observation, at least some members of this lineage can produce remarkable amounts of exopolymeric substances (Stehr et al., 1995b). The N. marina Lineage The N. marina lineage comprises three cultivated species, N. marina, N. aestuarii and Nitrosomonas sp. (represented by the isolates Nm 51 and Nm 63). All isolates originate from marine environments. Molecular signatures of this lineage have also been detected in marine samples (Stephen et al., 1996; Casciotti and Ward, 2001) as well as in enrichment cultures from such environments (Stephen et al., 1996). The presence of members of this lineage has been observed with molecular techniques in estuarine environments, coastal sand dunes and even in freshwater (aquaria) and a wastewater treatment plant (Kowalchuk et al., 1997; Speksnijder et al., 1998; Purkhold et al., 2000; Burrell et al., 2001; de Bie et al., 2001; Nicolaisen and Ramsing, 2002). Some authors, unfortunately, have lumped together the N. oligotropha lineage and the N. marina lineage (Kowalchuk and Stephen, 2001), although both lineages are phylogenetically distinct (Figs. 6 and 7) and the species of the two lineages show pronounced differences in ecophysiological characteristics and distribution patterns. The N. cryotolerans Lineage The N. cryotolerans lineage is represented by only one species, N. cryotolerans, and only one isolate exists in culture. This strain has been isolated from a marine environment and is obligatorily halophilic and urease positive. In molecular in situ analyses of AOB populations, N. cryotolerans has not yet been detected, although most of the commonly applied 16S rRNA gene primers do match this species. The Nm 143 Lineage Recently, a new lineage was identified within the genus Nitrosomonas (Purkhold et al., 2003; Fig. 7). This lineage has been provisionally named after Nitrosomonas sp. Nm143 and contains three additional isolates. It is not known whether these four isolates represent one or more than one species. All isolates were obtained from marine systems. Molecular signatures of this lineage were found in several marine and estuary systems (McCaig et al., 1994; Stephen et al., 1996; de Bie et al., 2001; Nicolaisen and Ramsing, 2002). Uncultured Lineage The so-called “environmental lineage 5” is the only major evolutionary lineage within the Nitrosomonas group which does not yet contain a cultured representative

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

(Stephen et al., 1996; Purkhold et al., 2000). The 16S rRNA sequences of this lineage were obtained from freshwater environments, marine systems and a coastal sand dune (McCaig et al., 1994; McCaig et al., 1999; Stephen et al., 1996; Kowalchuk et al., 1997; Speksnijder et al., 1998; Bano and Hollibaugh, 2000).

803

At least within the so-called “Nitrosospira cluster 3,” profound differences regarding physiology and distribution patterns exist because this cluster, in contrast to the other four clusters, comprises many morphologically as well ecophysiologically different species.

The Gammaproteobacterial AOB The Nitrosospira Lineage Members of the Nitrosospira lineage were postulated by some authors to be the dominant AOB in nature (Hiorns et al., 1995; Hastings et al., 1997; Ceccherini et al., 1998). However, this postulate is rather unsecured because it is based on findings obtained with nonquantitative PCRdependent methods often involving primers and probes not sufficiently covering some of the Nitrosomonas lineages (Tables 6 and 7). The Nitrosospira lineage represents a phylogenetically young group of AOB. Therefore, no highly reproducible phylogenetic substructure can be revealed within this group based on comparative 16S rRNA or amoA sequence analysis (see the section on Phylogeny of AOB in this Chapter). Nevertheless, a provisional subdivision system of this group has been suggested (Stephen et al., 1996; extended by Purkhold et al., 2000; Fig 8) and widely applied for interpretation of data from molecular AOB diversity surveys (for a review, see Kowalchuk and Stephen, 2001). However, it is questionable whether the suggested clustering defines units characterized by particular ecophysiological traits. Most isolates of the Nitrosospira lineage were obtained from terrestrial systems. Consistent with this finding, the current molecular dataset suggests that members of the Nitrosospira clusters 0, 2, 3 and 4 are predominantly found in terrestrial habitats (Fig. 12). Whether the reported occurrence of these clusters in freshwater habitats reflects that these bacteria are autochthonous freshwater inhabitants or soilborne organisms transferred to the water bodies cannot be decided. Nitrosospiras of cluster 1, for which no isolate exists, also occur in soil but are more frequently found in marine systems. The analyzed wastewater treatment plants rarely harbored nitrosospiras, and only members of clusters 2 and 3 were detected. Constructed wetlands might represent an exception to this rule, since mainly Nitrosospira -related 16S rRNA sequences were retrieved from a plant using rhizoremediation technology (Abd El Haleem et al., 2000). Nitrosospira strains Nsp57, 58 and 65, which were isolated from masonry and form two novel branches within this group (Fig. 8), are the only Nitrosospira isolates for which yet no closely related sequences were directly obtained from an environment.

The cultured strains of the two species of the genus Nitrosococcus, representing the gammaproteobacterial AOB, have been isolated from marine environments (all strains of N. oceani) and from a salt lagoon or from a salt lake (two isolates of N. halophilus), respectively. This habitat range is in accordance with the salt requirements and with the salt tolerances of the two species in laboratory experiments. Using immunological methods and molecular techniques, N. oceani was detected exclusively in marine environments but shown to be widely distributed (Ward and Perry, 1980; Ward, 1982; Ward and O’Mullan, 2002). Nitrosococcus halophilus has not yet been observed in nature via molecular techniques.

Dominant Populations of AOB in Different Environments Attempts to describe the dominant AOB populations of an environment should always take into careful consideration whether the retrieved species are autochthonous or allochthonous members of the analyzed ecosystem. Often species are introduced into environments from adjacent habitats. Since ecophysiological characteristics of distinct species of AOB often differ only minimally, introduced populations have a good chance to survive in the novel environment. Moreover, anthropogenic influence has created many new environments colonized by AOB that had no or only little time to adapt evolutionarily to these systems. In such cases, the sum of ecophysiological parameters will determine finally which of the species competing with each other will become the dominant representative.

Marine Environments A relatively clear situation is found in marine environments. In these ecosystems, generally those AOB are dominant which have a compatible salt requirement and a suitable salt tolerance. Consequently, the gammaproteobacterial AOB, N. oceani together with representatives of the Nitrosomonas marina lineage are the only AOB isolated with high frequency from marine environments. The second species of Nitrosococ-

804

H.-P. Koops et al.

cus, N. halophilus, seems to prefer environments with higher salt concentrations (salt lagoons or even salt lakes). Most of the above-mentioned organisms (except N. halophilus) can also be detected in marine systems using molecular methods (Stephen et al., 1996; Ward and O’Mullan, 2002). It is not clear why only one strain of N. cryotolerans, which also is ecophysiologically well adapted to these systems, has been isolated from marine samples. Members of the Nm143 lineage also seem to be distributed exclusively in marine environments. However, the ecophysiological properties of the latter group have not yet been investigated. Occasionally, species of the Nitrosomons europaea/Nc. mobilis lineage, owing to their relatively pronounced salt tolerance, were cultured from marine samples (Koops et al., 1976; Koops and Pommerening-Röser, 2001). Using quantitative dot blot, Hovanec and DeLong (1996) found that N. europaea -like bacteria were a major component of the microbial community in filters from seawater aquaria, but the primer (NSM1B) used for this analysis also has a full match to several members of the N. marina lineage (Tables 6 and 7). The 16S rRNA sequences of the Nitrosomonas environmental lineage 5 and Nitrosospira cluster 1, which both contain no cultured representatives, were frequently detected in marine environments (McCaig et al., 1994; McCaig et al., 1999; Stephen et al., 1996; Phillips et al., 1999; Bano and Hollibaugh, 2000; Hollibaugh et al., 2002; Fig. 12), but quantitative molecular data are missing. The retrieval of nitrosospiras in systems with elevated salt concentrations is surprising, since all cultured species of this lineage have a low salt tolerance in laboratory studies. The mere detection of 16S rRNA sequences of Nitrosospira cluster 1 in a marine environment does not prove that these organisms indeed possess a high salt tolerance and thrive in these systems. Alternatively, these nitrosospiras might be allochthonous inhabitants able to survive for extended periods in the ocean without significant physiological activity. It is thus important 1) to isolate and study a strain of this cluster and/or 2) to prove physiological activity of members of this cluster in situ for example by combination of FISH and microautoradiography (Lee et al., 1999).

Salt Lakes In salt lakes, the pH as well as the composition and concentration of salts vary significantly (Ward et al., 2000; Sorokin et al., 2001) thus creating different niches for AOB. Accordingly, different AOB were found in different salt lakes. From a salt lake in Saudi Arabia, a strain of

CHAPTER 3.2.12

Nitrosococcus halophilus was obtained (Koops et al., 1990), and from soda lakes located in Mongolia, alkali tolerant representatives of Nitrosomonas halophila have been isolated (Sorokin et al., 2001). In molecular analyses, N. europaea -like and N. eutropha -like organisms were detected in the hypersaline Mono Lake in California (Ward et al., 2000). However, the PCR primers used in this study (NitA and NitD) were not suited for a general AOB diversity study (Tables 6 and 7). To get a more general view of the AOB community composition in such environments, more investigations must be carried out.

Freshwater Environments In natural freshwater environments, members of the Nitrosomonas oligotropha lineage are generally the dominant AOB representatives. This is indicated by repeated isolation of members of this lineage from high MPN dilutions (Koops and Harms, 1985; Stehr et al., 1995a; Koops and Pommerening-Röser, 2001). Quantitative PCRindependent molecular data on AOB community composition in these systems are not available, but at least 16S rRNA sequences of this lineage were frequently detected in freshwater water bodies and sediments (Speksnijder et al., 1998; Radeva et al., 1999; Bollmann and Laanbroek, 2001; Whitby et al., 2001b; Regan et al., 2002) as well as in the Schelde estuary (Belgium and The Netherlands; de Bie et al., 2001). In addition, 16S rRNA sequences of Nitrosospira clusters 0, 2, 3 and 4 were harvested in molecular analyses of these habitats (Kowalchuk et al., 1998; Speksnijder et al., 1998; Whitby et al., 1999; Whitby et al., 2001b) and in a freshwater aquarium (Burrell et al., 2001). Moreover, using immunological methods, nitrosospiras were found in a eutrophic lake sediment (Smorzewski and Schmidt, 1991). However, these results do not necessarily indicate Nitrosospira dominance in freshwater systems because quantitative data are lacking, and several of the PCR primers applied in these studies do not sufficiently cover the N. oligotropha cluster (Purkhold et al., 2000; Tables 6 and 7). Occasionally, strains of N. europaea, N. eutropha and N. nitrosa have been isolated from eutrophicated rivers and eutrophic lakes (Koops and Pommerening-Röser, 2001). Furthermore, 16S rRNA sequences of N. europaea/Nc. mobilis, of Nitrosomonas environmental lineage 5 and of the N. marina lineage were retrieved from freshwater systems (Speksnijder et al., 1998; Whitby et al., 1999; Bollmann and Laanbroek, 2001; Burrell et al., 2001), but the significance of these findings remains unclear.

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

805

Artificial Aquatic Environments Urea and ammonia are the most frequently found nitrogen compounds in sewage. In wastewater treatment plants, AOB oxidize ammonia to nitrite, which is subsequently converted to nitrate by the nitrite-oxidizing bacteria. Nitrate is then removed from the sewage by denitrifying bacteria via anaerobic respiration. The slow growth rate of AOB and their susceptibility to pH- and temperature swings as well as to several sewage compounds is responsible for frequent failure of the nitrification in municipal and industrial wwtps. Isolation techniques indicate that in standard municipal wwtps, N. eutropha seems to be the dominant representative (Watson and Mandel, 1971b; Koops and Harms, 1985), but N. europaea and Nc. mobilis have also been repeatedly cultivated (Juretschko et al., 1998; Koops and Pommerening-Röser, 2001). All these species belong to the same Nitrosomonas lineage (Fig. 7) In industrial wwtps, the cultivation of representatives of members of the N. oligotropha lineage as well as N. nitrosa has been regularly reported (Koops and Harms, 1985; Suwa et al., 1997). In laboratory experiments, a remarkable high tolerance of members of the N. oligotropha lineage to heavy metals was observed, and the production of significant amounts of exopolymeric materials by these species was suggested to be the major reason for this tolerance (Stehr et al., 1995b). This resistance to heavy metals may be responsible for the presence of members of this lineage in special wwtps. During the last decade, molecular techniques were extensively applied to characterize AOB community structure and activity in activated sludge and biofilm from various wwtps. FISH revealed that AOB generally occur in tight microcolonies formed of up to several thousand individual cells (Fig. 14). These microcolonies cannot be easily dispersed using ultrasonic or Ultraturrax homogenizer treatments and thus contribute to the underestimation of AOB numbers in MPN assays. Typically, the AOB are detected in situ in close proximity to nitrite oxidizers reflecting the mutualistic relationship between both guilds (Mobarry et al., 1996; Schramm et al., 1996; Schramm et al., 1998; Juretschko et al., 1998; Okabe et al., 1999). Experiments using a combination of FISH with 14 C-bicarbonate microautoradiography demonstrated that almost all AOB microcolonies in activated sludge are physiologically active. Furthermore, AOB in wwtps can incorporate pyruvate (Daims et al., 2001a). Using quantitative FISH data, the specific activity of a single AOB cell in activated sludge has been estimated to be 2.3 ± 0.4 fmol of ammonia per hour (Daims

Fig. 14

Fig. 14. FISH detection of an AOB microcolony in a nitrifying biofilm from a sequencing batch biofilm reactor. AOB were stained by probe NEU, and the fluorescence signals were recorded by confocal laser scanning microscopy. Probeconferred fluorescence is shown in yellow, while autofluorescence of the biofilm is depicted in blue. Bar = 10 µm. The picture was kindly provided by Holger Daims.

et al., 2001c). According to FISH analyses, nitrosomonads are predominant over nitrosospiras in most activated sludge and biofilm systems (Mobarry et al., 1996; Schramm et al., 1996; Schramm et al., 2000; Juretschko et al., 1998; Okabe et al., 1999; Daims et al., 2001b; Gieseke et al., 2001). With the exception of Nitrosomonas cryotolerans, Nitrosomonas halophila and Nitrosomonas sp. Nm143, relatives of all other Nitrosomonas species occur in wwtps (Purkhold et al., 2000). Nitrosococcus mobilis, originally thought to be restricted to brackish water, is abundant in many wwtps (Juretschko et al., 1998; Daims et al., 2001b). Some nitrifying wwtps are dominated by a single AOB species (Juretschko et al., 1998), while other plants harbor a high diversity of AOB (Purkhold et al., 2000; Daims et al., 2001b; Gieseke et al., 2001). Whether and how differences in diversity of ammonia oxidizers are linked to the stability of the nitrification process is a yet unanswered question. Investigations of AOB populations in the raw sewage most probably also could give important insight into AOB diversity in sewage disposal plants.

Soils In acidic soils (heathlands, grasslands, forest soils, and marshy lands), where sufficient amounts of free ammonia are missing, ureasepositive species seem to be the exclusive representatives of the AOB. This has become obvious

806

H.-P. Koops et al.

via isolations as well as from molecular investigations (Kowalchuk and Stephen, 2001). Exclusively, species of the genera Nitrosospira and Nitrosovibrio could be detected. Molecular studies confirmed the predominance of nitrosospiras in acidic soils (Stephen et al., 1996; Smith et al., 2001) and indicated that, in particular, members of the Nitrosospira cluster 2 are well adapted to low pH terrestrial habitats (Kowalchuk et al., 1997; Stephen et al., 1998). A more detailed upto-date summary of these findings can be found in Kowalchuk and Stephen (2001). However, urease-positive strains of Nitrosolobus multiformis and of the genus Nitrosomonas, respectively, are missing in acidic soils. In more neutral soils, especially if agriculturally used, members of the N. communis lineage together with strains of Nitrosolobus multiformis are absolutely dominant in high MPN dilutions (MacDonald, 1986; Koops and PommereningRöser, 2001). In oligotrophic natural soils, especially if moderately acidic (pH around 6.0), strains of the N. oligotropha lineage as well as strains of the genera Nitrosospira or Nitrosovibrio have also been isolated (Koops and Pommerening-Röser, 2001).

Rocks and Building Stones AOB are also common on surfaces of rocks or building stones. Many strains have been isolated, and all were found to be members of the genera Nitrosospira or Nitrosovibrio (Spieck et al., 1992). Neither the closely related genus Nitrosolobus nor any of the lineages of the genus Nitrosomonas seem to be represented in these environments. Currently, the selective factors leading to these distribution patterns are speculative. Most likely, the acidic pH caused by the nitrification process is a crucial factor. Periodically appearing dryness or drastic change in temperature also may be selective for members of the genera Nitrosospira or Nitrosovibrio.

Literature Cited Aakra, A., J. B. Utaaker, I. F. Nes, and L. R. Bakken. 1999a. An evaluated improvement of the extinction dilution method for isolation of ammonia-oxidizing bacteria. J. Microbiol. Meth. 39:23–31. Aakra, A., J. B. Utaaker, and I. F. Nes. 1999b. RFLP of rRNA genes and sequencing of the 16S-23S rDNA intergenic spacer region of ammonia-oxidizing bacteria: A phylogenetic approach. Int. J. Syst. Bacteriol. 49(1):123–130. Aakra, A., M. Hesselsoe, and L. R. Bakken. 2000. Surface attachment of ammonia-oxidizing bacteria in soil. Microb. Ecol. 39:222–235. Aakra, A., J. B. Utaaker, and I. F. Nes. 2001a. Comparative phylogeny of the ammonia monooxygenase subunit A

CHAPTER 3.2.12 and 16S rRNA genes of ammonia-oxidizing bacteria. FEMS Microbiol. Lett. 205:237–242. Aakra, A., J. B. Utaaker, A. Pommerening-Roser, H.-P. Koops, and I. F. Nes. 2001b. Detailed phylogeny of ammonia-oxidizing bacteria determined by rDNA sequences and DNA homology values. Int. J. Syst. Evol. Microbiol. 51:2021–2030. Abd El Haleem, D., F. von Wintzingerode, A. Moter, H. Moawad, and U. B. Göbel. 2000. Phylogenetic analysis of rhizosphere associated β-subclass proteobacterial ammonia oxidizers in a municipal wastewater treatment plant based on rhizoremediation technology. Lett. Appl. Microbiol. 31:34–38. Abeliovich, A. 1987. Nitrifying bacteria in wastewater reservoirs. Appl. Environ. Microbiol. 53:754–760. Alzerreca, J. J., J. M. Norton, and M. G. Klotz. 1999. The amo operon in marine, ammonia–oxidizing gamma– proteobacteria. FEMS Microbiol. Lett. 180:21–29. Bano, N., and J. T. Hollibaugh. 2000. Diversity and distribution of DNA sequences with affinity to ammoniaoxidizing bacteria of the beta subdivision of the class Proteobacteria in the Arctic Ocean. Appl. Environ. Microbiol. 66:1960–1969. Baribeau, H., C. A. Kinner, J. R. Stephen, R. de Leon, P. A. Rochelle, and D. L. Clark. 2000. Presented at the American Water Works Association Water Quality Technology Conference Proceedings. Bartosch, S., I. Wolgast, E. Spieck, and E. Bock. 1999. Identification of nitrite-oxidizing bacteria with monoclonal antibodies recognizing the nitrite oxidoreductase. Appl. Environ. Microbiol. 65:4126–4133. Belser, L. W. 1979. Population ecology of nitrifying bacteria. Ann. Rev. Microbiol. 33:309–333. Belser, L. W., and E. L. Mays. 1982. Use of nitrifier activity measurements to estimate the efficiency of viable nitrifier counts in soilsand sediments. Appl. Environ. Microbiol. 43:945–948. Bollmann, A., and H. J. Laanbroek. 2001. Continuous culture enrichments of ammonia-oxidizing bacteria at low ammonium concentrations. FEMS Microbiol. Ecol. 37:211–221. Bruns, M. A., J. R. Stephen, G. A. Kowalchuk, J. I. Prosser, and E. A. Paul. 1999. Comparative diversity of ammonia oxidizer 16S rRNA gene sequences in native, tilled, and successional soils. Appl. Environ. Microbiol. 65:2994– 3000. Buchanan, R. E. 1917. Studies on the nomenclature and classification of bacteria. III: The families of the Eubacteriales. J. Bacteriol. 2:347–350. Burrell, P. C., C. M. Phalen, and T. A. Hovanec. 2001. Identification of bacteria responsible for ammonia oxidation in freshwater aquaria. Appl. Environ. Microbiol. 67:5791–5800. Burton, S. A., and J. I. Prosser. 2001. Autotrophic ammonia oxidation at low pH through urea hydrolysis. Appl. Environ. Microbiol. 67:2952–2957. Campbell, G. R. 2000. Community Analysis of Betasubgroup Ammonia-oxidising Bacteria in Sewage Sludge Amended Soil (PhD thesis). University of Aberdeen. Aberdeen, UK. Casciotti, K. L., and B. B. Ward. 2001. Dissimilatory nitrite reductase genes from autotrophic ammonia-oxidizing bacteria. Appl. Environ. Microbiol. 67:2213–2221. Ceccherini, M. T., M. Castialdini, C. Piovanelli, R. C. Hastings, A. J. McCarthy, M. Bazzicalupoet, and N. Miclaus. 1998. Effects of swine manure on autotrophic ammonia-

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

oxidizing bacteria in soil microcosms. Appl. Soil Ecol. 7:149–157. Chandler, D. P., R. W. Schreckhise, J. L. Smith, and H. Bolton. 1997. Electroelution to remove humic compounds from soil DNA and RNA extracts. J. Microbiol. Meth. 28:11– 19. Chang, Y. J., A. K. Hussain, J. R. Stephen, M. D. Mullen, D. C. White, and A. Peacock. 2001. Impact of herbizides on the abundance and structure of indigenous betasubgroup ammonia-oxidizer communities in soil microcosms. Environ. Toxicol. Chem. 20:2462–2468. Daims, H., J. L. Nielsen, P. H. Nielsen, K. H. Schleifer, and M. Wagner. 2001a. In situ characterization of Nitrospira-like nitrite-oxidizing bacteria active in wastewater treatment plants. Appl. Environ. Microbiol. 67:5273– 5284. Daims, H., U. Purkhold, L. Bjerrum, E. Arnold, P. A. Wilderer, and M. Wagner. 2001b. Nitrification in sequencing biofilm batch reactors: Lessons from molecular approaches. Water Sci. Tech. 43:9–18. Daims, H., N. B. Ramsing, K. H. Schleifer, and M. Wagner. 2001c. Cultivation-independent, semiautomatic determination of absolute bacterial cell numbers in environmental samples by fluorescence in situ hybridization. Appl. Environ. Microbiol. 67:5810–5818. de Bie, M. J. M., A. G. C. L. Speksnijder, G. A. Kowalchuk, T. Schuurmann, G. Zwart, J. R. Stephen, O. E. Diekmann, and H. J. Laanbroek. 2001. Shifts in the dominant populations of ammonia-oxidizing β-subclass Proeteobacteria along the eutrophic Schelde estuary. Aquat. Microb. Ecol. 23:225–236. De Boer, W., and H. J. Laanbroek. 1989. Ureolytic nitrification at low pH by Nitrosospira species. Arch. Microbiol. 152:178–181. De Boer, W., P. J. A. Klein-Gunnewiek, M. Veenhuis, E. Bock, and H. J. Laanbroek. 1991. Nitrification at low pH by aggregated autotrophic bacteria. Appl. Environ. Microbiol. 57:3600–3604. Dionisi, H. M., A. C. Layton, G. Harms, I. R. Gregory, K. G. Robinson, and G. S. Sayler. 2002. Quantification of Nitrosomonas oligotropha-like ammonia-oxidizing bacteria and Nitrospira spp. from full-scale wastewater treatment plants by competitive PCR. Appl. Environ. Microbiol. 68:245–253. Feray, C., B. Volat, V. V. Degrange, A. Clays-Josserand, and B. Montuelle. 1999. Assessment of three methods for detection and quantification of nitrite-oxidizing bacteria and nitrobacter in freshwater sediments (MPN-PCR, MPN-Griess, immunofluorescence). Microb. Ecol. 37:208–217. Gieseke, A., U. Purkhold, M. Wagner, R. Amann, and A. Schramm. 2001. Community structure and activity dynamics of nitrifying bacteria in a phosphate-removing biofilm. Appl. Environ. Microbiol. 67:1351–1362. Golovacheva, R. S. 1976. Thermophilic nitrifying bacteria from hot springs. Microbiology 45:329–331. Goreau, T. W., W. A. Kaplan, S. C. Wofsy, M. B. McElroy, F. W. Valois, and S. W. Watson. 1980. Production of NO2− and N2O by nitrifying bacteria at reduced concentrations of oxygen. Appl. Environ. Microbiol. 40:526–532. Guschin, D. Y., B. K. Mobarry, D. Proudnikov, D. A. Stahl, B. E. Rittmann, and A. D. Mirzabekov. 1997. Oligonucleotide microchips as genosensors for determinative and environmental studies in microbiology. Appl. Environ. Microbiol. 63:2397–2402.

807

Hall, G. H. 1986. Nitrification in lakes. In: J. I. Prosser (Ed.) Nitrification. IRL press. Washington, DC. 127–156. Harms, H., H.-P. Koops, and H. Wehrmann. 1976. An ammonia-oxidizing bacterium, Nitrosovibrio tenuis nov. gen. nov. sp. Arch. Microbiol. 108:105–111. Hastings, R. C., M. T. Ceccherini, N. Miclaus, J. R. Saunders, M. Bazzicalupo, and A. J. McCarthy. 1997. Direct molecular biological analysis of ammonia oxidising bacteria populations in cultivated soil plots treated with swine manure. FEMS Microbiol. Ecol. 23:45–54. Head, I. M., W. D. Hiorns, T. M. Embley, A. J. McCarthy, and J. R. Saunders. 1993. The phylogeny of autotrophic ammonia-oxidizing bacteria as determined by analysis of 16S ribosomal RNA gene sequences. J. Gen. Microbiol. 139:1147–1153. Helmer, C., S. Kunst, S. Juretschko, M. C. Schmid, K.-H. Schleifer, and M. Wagner. 1999. Nitrogen loss in a nitrifying biofilm system. Water Sci. Tech. 39:13–21. Henckel, T., M. Friedrich, and R. Conrad. 1999. Molecular analyses of the methane-oxidizing microbial community in rice field soil by targeting the genes of the 16S rRNA, particulate methane monooxygenase, and methanol dehydrogenase. Appl. Environ. Microbiol. 65:1980– 1990. Hermansson, A., and P. E. Lindgren. 2001. Quantification of ammonia-oxidizing bacteria in arable soil by real-time PCR. Appl. Environ. Microbiol. 67:972–976. Hiorns, W. D., R. C. Hastings, I. M. Head, A. J. McCarthy, J. R. Saunders, R. W. Pickup, and G. H. Hall. 1995. Amplification of 16S ribosomal RNA genes of autotrophic ammonia-oxidizing bacteria demonstrates the ubiquity of nitrosospiras in the environment. Microbiology 141:2793–800. Hollibaugh, J. T., N. Bano, and H. W. Ducklow. 2002. Widespread distribution in polar oceans of a 16S rRNA gene sequence with affinity to Nitrosospira-like ammoniaoxidizing bacteria. Appl. Environ. Microbiol. 68:1478– 1484. Holmes, A. J., A. Costello, M. E. Lidstrom, and J. C. Murrell. 1995. Evidence that particulate methane monooxygenase and ammonia monooxygenase may be evolutionarily related. FEMS Microbiol. Lett. 132:203–208. Holmes, A. J., P. Roslev, I. R. McDonald, N. Iversen, K. Henriksen, and J. C. Murrell. 1999. Characterization of methanotrophic bacterial populations in soils showing atmospheric methane uptake. Appl. Environ. Microbiol. 65:3312–3318. Hommes, N. G., L. A. Sayavedra-Soto, and D. J. Arp. 1998. Mutagenesis and expression of amo, which codes for ammonia monooxygenase in Nitrosomonas europaea. J. Bacteriol. 180:3353–3359. Horz, H. P., J. H. Rotthauwe, T. Lukow, and W. Liesack. 2000. Identification of major subgroups of ammonia-oxidizing bacteria in environmental samples by T-RFLP analysis of amoA PCR products. J. Microbiol. Meth. 39:197–204. Hoshino, T., N. Noda, S. Tsuneda, A. Hirata, and Y. Inamori. 2001. Direct detection by in situ PCR of the amoA gene in biofilm resulting from a nitrogen removal process. Appl. Environ. Microbiol. 67:5261–5266. Hovanec, T. A., and E. F. DeLong. 1996. Comparative analysis of nitrifying bacteria associated with freshwater and marine aquaria. Appl. Environ. Microbiol. 62:2888– 2896. Ivanova, I. A., J. R. Stephen, Y. J. Chang, J. Bruggemann, P. E. Long, J. P. McKinley, G. A. Kowalchuk, D. C. White, and S. J. Macnaughton. 2000. A survey of 16S rRNA and

808

H.-P. Koops et al.

amoA genes related to autotrophic ammonia-oxidizing bacteria of the beta-subdivision of the class Proteobacteria in contaminated groundwater. Can. J. Microbiol. 46:1012–1020. Jiang, Q. Q., and L. R. Bakken. 1999. Comparison of Nitrosospira strains isolated from terrestrial environments. FEMS Microbiol. Ecol. 30:171–186. Johnstone, B. H., and R. D. Jones. 1988. Physiological effects of long-term energy-source deprivation on the survival of a marine chemolithotrophic ammonia-oxidizing bacterium. Mar. Ecol. Prog. Ser. 49:295–303. Jones, R. D., R. Y. Morita, H.-P. Koops, and S. W. Watson. 1988. A new marine ammonium-oxidizing bacterium, Nitrosomonas cryotolerans sp. nov. Can. J. Microbiol. 34:1122–1128. Juretschko, S., G. Timmermann, M. Schmid, K. H. Schleifer, A. Pommerening-Roser, H. P. Koops, and M. Wagner. 1998. Combined molecular and conventional analyses of nitrifying bacterium diversity in activated sludge: Nitrosococcus mobilis and Nitrospira-like bacteria as dominant populations. Appl. Environ. Microbiol. 64:3042–3051. Klotz, M. G., and J. M. Norton. 1995. Sequence of an ammonia monooxygenase subunit A-encoding gene from Nitrosospira sp. NpAV. Gene 163:159–160. Koops, H.-P., H. Harms, and H. Wehrmann. 1976. Isolation of a moderate halophilic ammonia–oxidizing bacterium, Nitrosococcus mobilis nov. sp. Arch. Microbiol. 10:277– 282. Koops, H.-P., and H. Harms. 1985. Deoxyribonucleic acid homologies among 96 strains of ammonia-oxidizing bacteria. Arch. Microbiol. 141:214–218. Koops, H.-P., B. Böttcher, U. C. Möller, A. PommereningRöser, and G. Stehr. 1990. Description of a new species of Nitrosococcus. Arch. Microbiol. 154:244–248. Koops, H.-P., B. Böttcher, U. C. Möller, A. PommeringRöser, and G. Stehr. 1991. Classification of eight new species of ammonia-oxidizing bacteria: Nitrosomonas communis sp. nov., Nitrosomonas ureae sp. nov., Nitrosomonas aestuarii sp. nov., Nitrosomonas marina sp. nov., Nitrosomonas nitrosa sp. nov., Nitrosomonas eutropha sp. nov., Nitrosomonas oligotropha sp. nov., and Nitrosomonas halophila sp. nov. J. Gen. Microbiol. 137:1689–1699. Koops, H.-P., and U. C. Möller. 1992. The lithotrophic ammonia-oxidizing bacteria. In: A. Balows, H. G. Trü per, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York, NY. 2625–2637. Koops, H.-P., and A. Pommerening-Röser. 2001. Distribution and ecophysiology of the nitrifying bacteria emphasizing cultured species. FEMS Microbiol. Ecol. 37:1–9. Kowalchuk, G. A., J. R. Stephen, W. De Boer, J. I. Prosser, T. M. Embley, and J. W. Woldendorp. 1997. Analysis of ammonia-oxidizing bacteria of the beta subdivision of the class Proteobacteria in coastal sand dunes by denaturing gradient gel electrophoresis and sequencing of PCR-amplified 16S ribosomal DNA fragments. Appl. Environ. Microbiol. 63:1489–1497. Kowalchuk, G. A., Z. S. Naoumenko, P. J. L. Derikx, A. Felske, J. R. Stephen, and H. J. Laanbroek. 1998. Community analysis of ammonia-oxidising bacteria, in relation to oxygen availability in soils and root-oxygenated sediments, using PCR, DGGE and oligonucleotide probe hybridisation. FEMS Microbiol. Ecol. 27:339– 350.

CHAPTER 3.2.12 Kowalchuk, G. A., A. W. Stienstra, G. H. Heilig, J. R. Stephen, and J. W. Woldendorp. 2000a. Changes in the community structure of ammonia-oxidizing bacteria during secondary succession of calcareous grasslands. Environ. Microbiol. 2:99–110. Kowalchuk, G. A., A. W. Stienstra, G. H. Heilig, J. R. Stephen, and J. W. Woldendorp. 2000b. Molecular analysis of ammonia-oxidising bacteria in soil of successional grasslands of the Drentsche A (The Netherlands). FEMS Microbiol. Ecol. 31:207–215. Kowalchuk, G. A., and J. R. Stephen. 2001. Ammoniaoxidizing bacteria: A model for molecular microbial ecology. Ann. Rev. Microbiol. 55:485–529. Lee, N., P. H. Nielsen, K. H. Andreasen, S. Juretschko, J. L. Nielsen, K.-H. Schleifer, and M. Wagner. 1999. Combination of fluorescent in situ hybridization and microautoradiography: A new tool for structure-function analyses in microbial ecology. Appl. Environ. Microbiol. 65:1289–1297. Liebig, T., M. Wagner, L. Bjerrum, and M. Denecke. 2001. Nitrification performance and nitrifier community composition of a chemostat and a membrane-assisted bioreactor for the nitrification of sludge reject water. Bioproc. Biosyst. Engin. 24:203–210. Logemann, S., J. Schantl, S. Bijvank, M. van Loosdrecht, J. G. Kuenen, and M. Jetten. 1998. Molecular microbial diversity in a nitrifying reactor system without sludge retention. FEMS Microbiol. Ecol. 27:239–249. Ludwig, W., O. Strunk, S. Klugbauer, N. Klugbauer, M. Weizenegger, J. Neumaier, M. Bachleitner, and K.-H. Schleifer. 1998. Bacterial phylogeny based on comparative sequence analysis. Electrophoresis 19:554–568. MacDonald, R. M. 1986. Nitrification in soil: An introductory history. In: J. I. Prosser (Ed.) Nitrification. IRL Press. Oxford, UK. 1–16. McCaig, A. E., T. M. Embley, and J. I. Prosser. 1994. Molecular analysis of enrichment cultures of marine ammonia oxidisers. FEMS Microbiol. Lett. 120:363–367. McCaig, A. E., C. J. Phillips, J. R. Stephen, G. A. Kowalchuk, S. M. Harvey, R. A. Herbert, T. M. Embley, and J. I. Prosser. 1999. Nitrogen cycling and community structure of proteobacterial beta-subgroup ammonia-oxidizing bacteria within polluted marine fish farm sediments. Appl. Environ. Microbiol. 65:213–220. McTavish, H., J. A. Fuchs, and A. B. Hooper. 1993. Sequence of the gene coding for ammonia monooxygenase in Nitrosomonas europaea. J. Bacteriol. 175:2436–2444. Mendum, T. A., R. E. Sockett, and P. R. Hirsch. 1999. Use of molecular and isotopic techniques to monitor the response of autotrophic ammonia-oxidizing populations of the beta subdivision of the class Proteobacteria in arable soils to nitrogen fertilizer. Appl Environ. Microbiol. 65:4155–4162. Migula, W. 1900. System der Bakterien 2:194–195. Mobarry, B. K., M. Wagner, V. Urbain, B. E. Rittmann, and D. A. Stahl. 1996. Phylogenetic probes for analyzing abundance and spatial organization of nitrifying bacteria [published erratum appears in Appl. Environ. Microbiol.(1997), 63(2), p. 815]. Appl. Environ. Microbiol. 62:2156–2162. Morgenroth, E., A. Obermayer, E. Arnold, A. Brü hl, M. Wagner, and P. A. Wilderer. 2000. Effect of long-term idle periods on the performance of sequencing batch reactors. Water Sci. Tech. 41:105–113. Morris, S. A., S. Radajewski, T. W. Willison, and J. C. Murrell. 2002. Identification of the functionally active methan-

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

otroph population in a peat soil microcosm by stableisotope probing. Appl. Environ. Microbiol. 68:1446– 1453. Murray, R. G. E., and S. W. Watson. 1965. Structure of Nitrosocystis oceanus and comparison with Nitrosomonas and Nitrobacter. J. Bacteriol. 89:1594–1609. Nicolaisen, M. H., and N. B. Ramsing. 2002. Denaturing gradient gel electrophoresis (DGGE) approaches to study the diversity of ammonia-oxidizing bacteria. J. Microbiol. Meth. 50:189–203. Nogueira, R., L. F. Melo, U. Purkhold, S. Wuertz, and M. Wagner. 2002. Nitrifying and heterotrophic population dynamics in biofilm reactors: Effects of hydraulic retention time and the presence of organic carbon. Water Res. 36:469–481. Nold, S. C., J. Zhou, A. H. Devol, and J. M. Tiedje. 2000. Pacific Northwest marine sediments contain ammonia-oxidizing bacteria in the beta subdivision of the Proteobacteria. Appl. Environ. Microbiol. 66:4532– 4535. Norton, J. M., J. J. Alzerreca, Y. Suwa, and M. G. Klotz. 2002. Diversity of ammonia monooxygenase operon in autotrophic ammonia-oxidizing bacteria. Arch. Microbiol. 177:139–149. Okabe, S., H. Satoh, and Y. Watanabe. 1999. In situ analysis of nitrifying biofilms as determined by in situ hybridization and the use of microelectrodes. Appl. Environ. Microbiol. 65:3182–3191. Oved, T., A. Shaviv, T. Goldrath, R. T. Mandelbaum, and D. Minz. 2001. Influence of effluent irrigation on community composition and function of ammonia-oxidizing bacteria in soil. Appl. Environ. Microbiol. 67:3426– 3433. Painter, H. A. 1986. Nitrification in the treatment of sewage and waste waters. In: J. I. Prosser (Ed.) Nitrification. IRL Press. Oxford, UK. 185–211. Pedersen, K., J. Arlinger, L. Hallback, and C. Petersson. 1996. Investigations of subterranean bacteria in deep crystalline bedrock and their importance for the disposal of nuclear waste. Can. J. Microbiol. 42:382–391. Phillips, C. J., Z. Smith, T. M. Embley, and J. I. Prosser. 1999. Phylogenetic differences between particle-associated and planktonic ammonia-oxidizing bacteria of the beta subdivision of the class Proteobacteria in the Northwestern Mediterranean Sea. Appl. Environ. Microbiol. 65:779–786. Phillips, C. J., D. Harris, S. L. Dollhopf, K. L. Gross, J. I. Prosser, and E. A. Paul. 2000. Effects of agronomic treatments on structure and function of ammoniaoxidizing communities. Appl. Environ. Microbiol. 66:5410–5408. Pinck, C., C. Coeur, P. Potier, and E. Bock. 2001. Polyclonal antibodies recognizing the AmoB protein of ammonia oxidizers of the beta-subclass of the class Proteobacteria. Appl. Environ. Microbiol. 67:118–124. Pommerening-Röser, A. 1993. Untersuchungen zur Phylogenie Ammoniak oxidierender Bakterien (PhD thesis). University of Hamburg. Hamburg, Germany. Pommerening-Röser, A., G. Rath, and H.-P. Koops. 1996. Phylogenetic diversity within the genus Nitrosomonas. Syst. Appl. Microbiol. 19:344–351. Princic, A., I. I. Mahne, F. Megusar, E. A. Paul, and J. M. Tiedje. 1998. Effects of pH and oxygen and ammonium concentrations on the community structure of nitrifying bacteria from wastewater. Appl. Environ. Microbiol. 64:3584–3590.

809

Purkhold, U., A. Pommerening-Röser, S. Juretschko, M. C. Schmid, H.-P. Koops, and M. Wagner. 2000. Phylogeny of all recognized species of ammonia oxidizers based on comparative 16S rRNA and amoA sequence analysis: Implications for molecular diversity surveys. Appl. Environ. Microbiol. 66:5368–5382. Purkhold, U., M. Wagner, G. Timmermann, A. PommereningRöser, and H.-P. Koops. 2003. 16S rRNA and amoAbased phylogeny of 12 novel betaproteobacterial ammonia oxidizing isolates: Extension of the data set and proposal of a new lineage within the Nitrosomonads. Int. J. Syst. Evol. Microbiol. 53:1485–1494. Radajewski, S., G. Webster, D. S. Reay, S. A. Morris, P. Ineson, D. B. Nedwell, J. I. Prosser, and J. C. Murrell. 2002. Identification of active methylotroph populations in an acidic forest soil by stable-isotope probing. Microbiology 148:2331–2342. Radeva, G., K. Flemming, and S. Selenska–Pobell. 1999. Molecular analysis of bacterial populations in ground water pollutes with heavy metals. In: G. Bernhard (Ed.) Annual Report, Institute of Biochemistry. Institute of Biochemistry. 57. Rath, G. 1996. PhD thesis. University of Hamburg. Hamburg, Germany. Regan, J. M., G. W. Harrington, and D. R. Noguera. 2002. Ammonia- and nitrite-oxidizing bacterial communities in a pilot-scale chloraminated drinking water distribution system. Appl. Environ. Microbiol. 68:73–81. Rotthauwe, J. H., W. de Boer, and W. Liesack. 1995. Comparative analysis of gene sequences encoding ammonia monooxygenase of Nitrosospira sp. AHB1 and Nitrosolobus multiformis C-71. FEMS Microbiol. Lett. 133:131– 135. Rotthauwe, J. H., K. P. Witzel, and W. Liesack. 1997. The ammonia monooxygenase structural gene amoA as a functional marker: molecular fine-scale analysis of natural ammonia-oxidizing populations. Appl. Environ. Microbiol. 63:4704–4712. Sakano, Y., K. D. Pickering, P. F. Strom, and L. J. Kerkhof. 2002. Spatial distribution of total, ammonia-oxidizing, and denitrifying bacteria in biological wastewater treatment reactors for bioregenerative life support. Appl. Environ. Microbiol. 68:2285–2293. Schmid, M., U. Twachtmann, M. Klein, M. Strous, S. Juretschko, M. Jetten, J. W. Metzger, K.-H. Schleifer, and M. Wagner. 2000. Molecular evidence for a genuslevel diversity of bacteria capable of catalyzing anaerobic ammonium oxidation. Syst. Appl. Microbiol. 23:93– 106. Schmidt, I., and E. Bock. 1997. Anaerobic ammonia oxidation with nitrogen dioxide by Nitrosomonas eutropha. Arch. Microbiol. 167:106–111. Schmidt, I., and E. Bock. 1998. Anaerobic ammonia oxidation with nitrogen dioxide by cell-free extracts of Nitrosomonas eutropha. Ant. v. Leeuwenhoek 73:271–278. Schramm, A., L. H. Larsen, N. P. Revsbech, N. B. Ramsing, R. Amann, and K.-H. Schleifer. 1996. Structure and function of a nitrifying biofilm as determined by in situ hybridization and the use of microelectrodes. Appl. Environ. Microbiol. 62:4641–4647. Schramm, A., D. De Beer, M. Wagner, and R. Amann. 1998. Identification and activities in situ of Nitrosospira and Nitrospira spp. as dominant populations in a nitrifying fluidized bed reactor. Appl. Environ. Microbiol. 64:3480–5.

810

H.-P. Koops et al.

Schramm, A., D. de Beer, J. C. van den Heuvel, S. Ottengraf, and R. Amann. 1999. Microscale distribution of populations and activities of Nitrosospira and Nitrospira spp. along a macroscale gradient in a nitrifying bioreactor: quantification by in situ hybridization and the use of microsensors. Appl. Environ. Microbiol. 65:3690–3696. Schramm, A., D. De Beer, A. Gieseke, and R. Amann. 2000. Microenvironments and distribution of nitrifying bacteria in a membrane-bound biofilm. Environ. Microbiol. 2:680–6. Silyn-Roberts, G., and G. Lewis. 2001. In situ analysis of Nitrosomonas spp. in wastewater treatment wetland biofilms. Water Res. 35:2731–2739. Sinigalliano, C. D., D. N. Kuhn, and R. D. Jones. 1995. Amplification of the amoA gene from diverse species of ammonium-oxidizing bacteria and from an indigenous bacterial population from seawater. Appl. Environ. Microbiol. 61:2702–2706. Smith, Z., A. E. McCaig, J. R. Stephen, T. M. Embley, and J. I. Prosser. 2001. Species diversity of uncultured and cultured populations of soil and marine ammonia oxidizing bacteria. Microb. Ecol. 42:228–237. Smorzewski, W. T., and E. I. Schmidt. 1991. Numbers, activities, and diversity of autotrophic ammonia-oxidizing bacteria in a freshwater, eutrophic lake sediment. Can. J. Microbiol. 37:828–833. Sorokin, D. Y., G. Muyzer, T. Brinkhoff, J. G. Kuenen, and M. S. M. Jetten. 1998. Isolation and characterization of a novel facultatively alkaliphilic Nitrobacter species, N. alkalicus sp. nov. Arch. Microbiol. 170:345– 352. Sorokin, D., T. Tourova, M. C. Schmid, M. Wagner, H. P. Koops, J. G. Kuenen, and M. Jetten. 2001. Isolation and properties of obligately chemolithoautotrophic and extremely alkali-tolerant ammonia-oxidizing bacteria from Mongolian soda lakes. Arch. Microbiol. 176:170– 177. Speksnijder, A. G., G. A. Kowalchuk, K. Roest, and H. J. Laanbroek. 1998. Recovery of a Nitrosomonas-like 16S rDNA sequence group from freshwater habitats. Syst. Appl. Microbiol. 21:321–330. Speksnijder, A. G., G. A. Kowalchuk, S. De Jong, E. Kline, J. R. Stephen, and H. J. Laanbroek. 2001. Microvariation artifacts introduced by PCR and cloning of closely related 16S rRNA gene sequence. Appl. Environ. Microbiol. 67:469–472. Spieck, E., M. Meincke, and E. Bock. 1992. Taxonomic diversity of Nitrosovibrio strains isolated from building stones. FEMS Microbiol. Ecol. 102:21–26. Stackebrandt, E., R. G. E. Murray, and H. G. Trü per. 1988. Proteobacteria classis nov., a name for the phylogenetic taxon that includes the “purple bacteria and their relatives”. Int. J. Syst. Bacteriol. 38:321–325. Stackebrandt, E., and B. M. Goebel. 1994. Taxonomic note: a place for DNA-DNA reassociation and 16S rRNA sequence analysis in the present species definition in bacteriology. Int. J. Syst. Bacteriol. 44:846–849. Stehr, G., B. Böttcher, P. Dittberner, G. Rath, and H. P. Koops. 1995a. The ammonia-oxidizing nitrifying population of the River Elbe estuary. FEMS Microbiol. Ecol. 17:177–186. Stehr, G., S. Zörner, B. Böttcher, and H.-P. Koops. 1995b. Exopolymers: An ecological characteristic of a flocattached, ammonia-oxidizing bacterium. Microb. Ecol. 30:115–126.

CHAPTER 3.2.12 Stephen, J. R., A. E. McCaig, Z. Smith, J. I. Prosser, and T. M. Embley. 1996. Molecular diversity of soil and marine 16S rRNA gene sequences related to β-subgroup ammonia-oxidizing bacteria. Appl. Environ. Microbiol. 62:4147–4154. Stephen, J. R., G. A. Kowalchuk, M. A. V. Bruns, A. E. McCaig, C. J. Phillips, T. M. Embley, and J. I. Prosser. 1998. Analysis of beta-subgroup proteobacterial ammonia oxidizer populations in soil by denaturing gradient gel electrophoresis analysis and hierarchical phylogenetic probing. Appl. Environ. Microbiol. 64:2958–2965. Stephen, J. R., Y. J. Chang, S. J. Macnaughton, G. A. Kowalchuk, K. T. Leung, C. A. Flemming, and D. C. White. 1999. Effect of toxic metals on indigenous soil betasubgroup proteobacterium ammonia oxidizer community structure and protection against toxicity by inoculated metal-resistant bacteria. Appl. Environ. Microbiol. 65:95–101. Suwa, Y., Y. Imamura, T. Suzuki, T. Tashiro, and Y. Urushigawa. 1994. Ammonia-oxidizing bacteria with different sensitivities to (NH4 )2 SO4 in activated sludge. Water Res. 28:1523–1532. Suwa, Y., T. Sumino, and K. Noto. 1997. Phylogenetic relationships of activated sludge isolates of ammonia oxidizers with different sensitivities to ammonium sulfate. J. Gen. Appl. Microbiol. 43:373–379. Szwerinski, H., S. Gaiser, and D. Bardtke. 1985. Immunofluorescence for the quantitative determination of nitrifying bacteria: Interference of the test in biofilm reactors. Appl. Microbiol. Biotechnol. 21:125–128. Takahashi, R., N. Kondo, K. Usui, T. Kanehira, M. Shinohara, and T. Tokuyama. 1992. Pure isolation of a new chemoautotrophic ammonia-oxidizing bacterium on a gellan gum plate. J. Ferment. Bioeng. 74:52–54. Teske, A., E. Alm, J. M. Regan, T. S., B. E. Rittmann, and D. A. Stahl. 1994. Evolutionary relationships among ammonia- and nitrite-oxidizing bacteria. J. Bacteriol. 176:6623–6630. Tokuyama, T., N. Yoshida, T. Matasuishi, N. Takahashi, T. Takahashi, T. Kanehira, and M. Shinohara. 1997. A new psychrotrophic ammonia oxidizing bacterium Nitrosovibrio sp. TYM9. J. Ferment. Bioeng. 83:377–380. Trü per, H. G., and L. de Clari. 1997. Taxonomic note: Necessary correction of specific epithets formed as substantives (nouns) “in apposition”. Int. J. Syst. Bacteriol. 47:908–909. Utaaker, J. B., L. Bakken, Q. Q. Jiang, and I. F. Nes. 1995. Phylogenetic analysis of seven new isolates of ammoniaoxidizing bacteria based on 16S rRNA gene sequences. Syst. Appl. Microbiol. 18:549–559. Utaaker, J. B., and I. F. Nes. 1998. A qualitative evaluation of the published oligonucleotides specific for the 16S rRNA gene sequences of the ammonia-oxidizing bacteria. Syst. Appl. Microbiol. 21:72–88. Völsch, A., W. F. Nader, H. K. Geiss, G. Nebe, and C. Birr. 1990. Detection and analysis of two serotypes of ammonia-oxidizing bacteria in sewage plants by flow cytometry. Appl. Environ. Microbiol. 56:2430–2435. Voytek, M. A., and B. B. Ward. 1995. Detection of ammonium-oxidizing bacteria of the beta-subclass of the class Proteobacteria in aquatic samples with the PCR. Appl. Environ. Microbiol. 61:1444–1450. Voytek, M. A. 1996. Relative Abundance and Species Diversity of Autotrophic Ammonia Oxidizing Bacteria in Aquatic Systems (PhD thesis). University of California at Santa Cruz. Reston, Virginia.

CHAPTER 3.2.12

The Lithoautotrophic Ammonia-Oxidizing Bacteria

Wagner, M., G. Rath, R. Amann, H.-P. Koops, and K.-H. Schleifer. 1995. In situ identification of ammoniaoxidizing bacteria. Syst. Appl. Microbiol. 18:251–264. Ward, B. B., and M. J. Perry. 1980. Immunofluoreszent assay for the marine ammonium-oxidizing bacterium Nitrosomonas oceanus. Appl. Environ. Microbiol 39:913–918. Ward, B. B. 1982. Oceanic distribution of ammoniumoxidizing bacteria determined by immunofluorscent assay. J. Mar. Res. 40:1155–1172. Ward, B. B., and A. F. Carlucci. 1985. Marine ammonia- and nitrit-oxidizing bacteria: serological diversity determined by immunofluorescence in sewage plants by flow cytometry. Appl. Environ. Microbiol. 50:194–201. Ward, B. B., M. A. Voytek, and K. Witzel. 1997. Phylogenetic diversity of natural populations of ammonia oxidizers investigated by specific PCR amplification. Microb. Ecol. 33:87–96. Ward, B. B., D. P. Martino, M. C. Diaz, and S. B. Joye. 2000. Analysis of ammonia-oxidizing bacteria from hypersaline Mono Lake, California, on the basis of 16S rRNA sequences. Appl. Environ. Microbiol. 66:2873– 2881. Ward, B. B., and G. D. O’Mullan. 2002. Worldwide distribution of Nitrosococcus oceani, a marine ammoniaoxidizing gamma-proteobacterium, detected by PCR and sequencing of 16S rRNA and amoA genes. Appl. Environ. Microbiol. 68:4153–4157. Watson, S. W. 1965. Characteristics of a marine nitrifying bacterium, Nitrosocystis oceanus sp. n. Limnol. Oceanogr. 10 (Suppl.):R274–R289. Watson, S. W., and C. C. Remsen. 1970. Cell envelope of Nitrosocystis oceanus. J. Ultrastr. Res. 33:148–160. Watson, S. W. 1971a. Taxonomic considerations of the family Nitrobacteraceae Buchanan: Requests for opinions. Int. J. Syst. Bacteriol. 21:254–270. Watson, S. W., and M. Mandel. 1971b. Comparison of the morphology and deoxyribonucleid acid composition of 27 strains of nitrifying bacteria. J. Bacteriol. 107:563– 569. Watson, S. W., L. B. Graham, C. C. Remsen, and F. W. Valois. 1971c. A lobular, ammonia oxidizing bacterium, Nitrosolobus multiformis, nov. sp. Arch. Microbiol. 76:183–203. Watson, S. W., F. W. Valois, and J. B. Waterbury. 1981. The family Nitrobacteraceae. In: M. P. Starr, H. Stolp, and H. Trü per (Eds.) The Prokaryotes. Springer-Verlag. Berlin, Germany. 1:1005–1022. Watson, S. W., E. Bock, H. Harms, H.-P. Koops, and A. B. Hooper. 1989. Nitrifying bacteria. In: J. T. Staley, M. P. Bryant, N. Pfennig, and J. G. Holt (Eds.) Bergey’s Manual of Systematic Bacteriology. Williams and Wilkins. Baltimore, MD. 3:1808–1834. Webster, G., T. M. Embley, and J. I. Prosser. 2002. Grassland management regimens reduce small-scale heterogeneity and species diversity of beta-proteobacterial ammonia oxidizer populations. Appl. Environ. Microbiol. 68:20– 30.

811

Whitby, C. B., J. R. Saunders, J. Rodriguez, R. W. Pickup, and A. McCarthy. 1999. Phylogenetic differentiation of two closely related Nitrosomonas spp. that inhabit different sediment environments in an oligotrophic freshwater lake. Appl. Environ. Microbiol. 65:4855– 4862. Whitby, C. B., G. Hall, R. Pickup, J. R. Saunders, P. Ineson, N. R. Parekh, and A. McCarthy. 2001a. 13C incorporation into DNA as a means of identifying the active components of ammonia-oxidizer populations. Lett. Appl. Microbiol. 32:398–401. Whitby, C. B., J. R. Saunders, R. W. Pickup, and A. J. McCarthy. 2001b. A comparison of ammonia-oxidiser populations in eutrophic and oligotrophic basins of a large freshwater lake. Ant. v. Leeuwenhoek 79:179–188. Wilhelm, R., A. Abeliovich, and A. Nejidat. 1998. Effect of long-term ammonia starvation on the oxidation of ammonia and hydroxylamine by Nitrosomonas europaea. J. Biochem. 124:811–815. Winogradsky, S. 1890. Recherches sur les organismes de la nitrification. Ann. Inst. Pasteur 4:213–331. Winogradsky, S. 1892. Contributions a la morphologie des organismes de la nitrification. Arch. Sci. Biol. (St. Petersburg) 1:88–137. Winogradsky, S., and H. Winogradsky. 1933. Etudes sur la microbiologie du sol. VII: Nouvelles recherches sur les organismes de la nitrification. Ann. Inst. Pasteur 50. Winogradsky, H. 1937. Contribution a l’étude de la microflore nitrificatrice des boues activées de Paris. Ann. Inst. Pasteur 58:326–340. Wintzigerode, F., and U. B. Goebel. 1997. Determination of microbial diversity in environmental samples: Pitfalls of PCR-based analysis. FEMS Microbiol. Rev. 21:213– 229. Woese, C. R., W. G. Weisburg, B. J. Paster, C. M. Hahn, R. S. Tanner, N. R. Krieg, H.-P. Koops, H. Harms, and E. Stackebrandt. 1984. The phylogeny of the purple bacteria: the beta subdivision. Syst. Appl. Microbiol. 5:327– 336. Woese, C. R., W. G. Weisburg, C. M. Hahn, B. J. Paster, L. B. Zablen, B. J. Lewis, T. J. Macke, W. Ludwig, and E. Stackebrandt. 1985. The phylogeny of the purple bacteria: The gamma subdivision. Syst. Appl. Microbiol. 6:25– 33. Wu, L., D. K. Thompson, G. Li, R. A. Hurt, J. M. Tiedje, and J. Zhou. 2001. Development and evaluation of functional gene arrays for detection of selected genes in the environment. Appl. Environ. Microbiol. 67:5780– 5790. Yamagata, A., J. Kato, R. Hirota, A. Kuroda, T. Ikeda, N. Takiguchi, and H. Ohtake. 1999. Isolation and characterization of two cryptic plasmids in the ammoniaoxidizing bacterium Nitrosomonas sp. strain ENI-11. J. Bacteriol. 181:3375–3381. Zart, D., I. Schmidt, and E. Bock. 2000. Significance of gaseous NO for ammonia oxidation by Nitrosomonas eutropha. Ant. v. Leeuwenhoek 77:49–55.

Prokaryotes (2006) 5:812–827 DOI: 10.1007/0-387-30745-1_37

CHAPTER 3.2.13 ehT

suneG

su l l i cabo i hT

The Genus Thiobacillus LESLEY A. ROBERTSON AND J. GIJS KUENEN

Introduction

Habitats

Ever since the genus Thiobacillus was first described in 1904, the ability to grow while using a reduced sulfur compound as a source of energy has been considered sufficiently important taxonomically to merit classifying all Gram-negative, sulfur-oxidizing, nonphototrophic rods in this genus. However, as studies using modern taxonomic methods began to reveal that some of the species are only superficially related, virtually every paper describing a new Thiobacillus species in the last decade has mentioned the need to reorganize the genus. Most of this reorganization has now been done (Kelly and Wood, 2000a), and the authors are grateful to D.P. Kelly and A. Wood for permission to see and use their resulting manuscripts prior to publication. The effect of the reorganization is dramatic (Table 1). In the last edition of The Prokaryotes, the genus Thiobacillus included 17 species. There are now only three species left—all autotrophs, and all members of the β-subclass of the Proteobacteria. The 4 or 5 facultative autotrophs that fall into the β-subclass have now been assigned to a new genus—Thiomonas. Most of the other former Thiobacillus species fall into the γ-subclass, together with the members of the genus Thiomicrospira. This includes both the well-known acidiphilic and recently discovered alkaliphilic “Thiobacillus” species, as well as the marine species. However, because the reorganization is still so new and most researchers will look for these organisms in one group, many of them will be dealt with in this chapter, with the exception of the γ group. Thiobacillus acidophilus and Thiobacillus versutus have both been transferred to existing genera (see Table 1). Similarly, Thiosphaera pantotropha has now been reclassified as Paracoccus pantotrophus and will be described in the chapter on that genus. The last member of the γ group, Thiobacillus novellus, is now Starkeya novella. Within this chapter, the generic terms “thiobacilli” and “colorless sulfur bacteria” refer to the whole group, regardless of their new classification.

In nature, the distribution of reduced inorganic sulfur compounds is only one of the factors governing the presence and activity of the colorless sulfur bacteria, and a complex of chemical, physical, and microbial interactions is almost as important a controlling force. One crucial factor is the requirement for the simultaneous presence of an electron donor (the reduced sulfur compound) and electron acceptor (i.e., oxygen or nitrogen oxides). Sulfide is one of the most important natural substrates, and many of the colorless sulfur bacteria grow in the narrow zones and gradients where sulfide and oxygen coexist (Nelson and Jannasch, 1983). Such gradients can be found, for example, in stratified lakes and at the interface between aerobic water and an anaerobic sediment, but may be even more common in microniches in environments that contain anaerobic pockets (e.g., sediments and wet soils). It must be realized that extremely high turnover rates of sulfide can be observed at very low levels of sulfide and oxygen (3,000 Da) hemolytic activity was described in one strain of B. cepacia that was able to induce nucleosomal degradation (consistent with apoptosis) in human neutrophils and in a mouse macrophage cell line (Hutchison et al., 1998). There does not appear to be any correlation between production of extracellular virulence factors and transmissibility of epidemic B. cepacia isolates. Some strains of the B. cepacia complex have been shown to invade and survive in cultured epithelial cells and macrophages (Burns et al., 1996; Martin and Mohr, 2000; Saini et al., 1999). Burkholderia cepacia also has been reported to be resistant to growth inhibition by epithelial cell β-defensins and resistant to nonoxidative neutrophil-killing mechanisms compared to P. aeruginosa (Baird et al., 1999; Speert et al., 1994). The ability to acquire iron from host sources is a prerequisite for the successful establishment and maintenance of most bacterial infections. Yang et al. have demonstrated that 84 : 84 B. pseudomallei strains examined during their studies tested positive for siderophore production using the chrome azurol S (CAS) assay. A structural and chemical analysis of the siderophore synthesized by B. pseudomallei U7 confirmed the molecule was approximately 1,000 Da in size, was water soluble with a yellow-green fluorescence, and belonged to the hydroxamate class (Yang et al., 1991). Furthermore, studies also have demonstrated the siderophore was capable of scavenging iron from both lactoferrin and

CHAPTER 3.2.16

transferrin in vitro (Yang et al., 1993). The name malleobactin has been proposed for this compound (Yang et al., 1991). Although B. pseudomallei isolates can express an impressive array of both secreted and cellassociated antigens, the role(s) of these products in the pathogenesis of disease to date have been relatively ill defined. One of the primary reasons for this has been the lack of suitable techniques for genetically manipulating the organism. Owing to the recent application of a transposon-based mutagenesis system for use in B. pseudomalle, genetic loci, which encode a number of these putative virulence determinants and protective antigens, have been identified and characterized (DeShazer et al., 1997; DeShazer et al., 1998). It has been shown previously that B. pseudomallei isolates can secrete antigens that demonstrate biological activities consistent with proteases, lecithinases, lipases and hemolysins (Esselman and Liu, 1961; Ashdown and Koehler, 1990; Sexton et al., 1994). However, though the importance of these factors has been implicated in the pathogenesis of the disease, only the protease has been characterized to date. Studies conducted by Sexton et al. (1994) have confirmed the presence of a 36-kDa antigen with associated proteolytic activities in B. pseudomallei culture supernatants. In particular, a protease expressed by B. pseudomallei (isolate 319a) was found to be a metalloenzyme requiring iron for maximal protease activity and to be optimally active at pH 8.0 and 60°C (Sexton et al., 1994). Furthermore, monoclonal antibodies (MAb) raised against a Pseudomonas aeruginosa alkaline protease were cross-reactive with this antigen (Sexton et al., 1994). Most recently, an 11.8-kb chromosomal locus in B. pseudomallei has been identified that demonstrates a high degree of homology to operons that encode for the products of the main terminal branch of the general secretory pathway (GSP; Pugsley, 1993). Further characterization of the open-reading frames in this locus have confirmed that their orientation and physical arrangement are virtually identical to the pul gene cluster of Klebsiella oxytoca (Pugsley, 1993). Not surprisingly, the phenotypic analysis of individual transposon mutants also has confirmed their inability to secrete antigens associated with protease, lipase and lecithinase into the extracellular milieu. Interestingly, although the protease, lipase and lecithinase may play a small role in the pathogenesis of acute melioidosis, mutants deficient in their ability to secrete these particular exoenzymes were not severely attenuated in their ability to cause a fulminating illness (DeShazer et al., 1999). In the mid-1950s, several studies demonstrated that filter-sterilized B. pseudomallei culture

CHAPTER 3.2.16

supernatants were lethal for mice and hamsters when administered parenterally (Nigg et al., 1955; Heckly and Nigg, 1958; Heckly, 1964). These results were consistent with the fulminating illnesses observed in animals following inoculation with viable bacteria and suggested that B. pseudomallei strains might be capable of secreting a lethal toxin. In studies conducted by Ismail et al. (1987), mouse lethal, thermolabile toxin was presumably purified to homogeneity and characterized as a 31-kDa protein. Haase et al. (1997) have also described the presence of cytotoxic activity in culture filtrates. Their results, however, suggest that the antigen is only 3 kDa in size and that the cytotoxic activity in this instance most likely is due to the presence of a small peptide. Recently it has been reported that a rhamnolipid purified from B. pseudomallei culture supernatants demonstrates a cytotoxic effect against HL60 and HeLa cell lines (Haubler et al., 1998). Because this activity can be neutralized by albumin, however, it is unlikely to be of consequence in the pathogenesis of B. pseudomallei infections. These results have not been reproduced in animal models, even when using preparations concentrated by lyophilization (Brett et al., 1997; Brett et al., 1998). Burkholderia pseudomallei strains can synthesize capsular antigens (Smith et al., 1987; Leelarasamee and Bovornkitti, 1989), and they may play an important role in the pathogenesis of melioidosis. Whereas in vitro studies have determined that encapsulated B. pseudomallei strains are as susceptible to phagocytic uptake by polymorphonuclear leukocytes (PMNs) as non-encapsulated variants, evidence tends to suggest that the presence of exopolysaccharide confers upon them the ability to resist the bactericidal effects of the phagolysosomal environment (Smith et al., 1987; Pruksachartvuthi et al., 1990). This feature of B. pseudomallei strains may help to explain why these organisms are able to remain latent in a host for as long as 26 years. Recently, Steinmetz et al. isolated and purified a high-molecular-weight capsular antigen (150 kDa) from B. pseudomallei NCTC 7431 and succeeded in raising a Mab against it (Steinmetz et al., 1995). They were able to demonstrate the reactivity of both mucoid and non-mucoid strains with the MAb, suggesting that the capsular antigen is constitutively expressed by B. pseudomallei strains. Interestingly, temperature appeared to have little effect on the synthesis of the exopolysaccharide inasmuch as B. pseudomallei strains grown at both 15 and 37°C were Mab-reactive. Furthermore, an assay utilizing a variety of Pseudomonas spp. and Burkholderia spp. as controls was able to

The Genus Burkholderia

855

confirm the specificity of the MAb for B. pseudomallei and B. mallei strains only (Steinmetz et al., 1995). More recently, Masoud et al. (1997) have been successful at elucidating the chemical and structural characteristics of a capsular polysaccharide isolated from the virulent clinical isolate B. pseudomallei 304b. Their results demonstrated that the exopolysaccharide was a linear unbranched polymer of repeating tetrasaccharide units having the structure (-3)-2-O-Ac-β-D-Galp-(1-4)-α-D-Galp-(1-3)-β-DGalp-(1-5)-β-D-KDOp-(2-) (where KDO is 3deoxy-D-manno-2-octulosonic acid). Similarly, Nimtz et al. (1997) have demonstrated that a structurally identical capsular antigen is expressed by B. pseudomallei NCTC 7431. Studies by both groups have also shown that patient sera reacted strongly with the purified carbohydrate antigens, indicating that this carbohydrate polymer is most likely expressed in vivo (Steinmetz et al., 1995; Masoud et al., 1997). Previous studies have confirmed that the lipopolysaccharide (LPS) antigens expressed by B. pseudomallei strains are highly conserved throughout this species (Pitt et al., 1992). In fact, serological evidence suggests only one serotype of B. pseudomallei (Bryan et al., 1994). To investigate this phenomenon, Perry et al. have characterized the LPS antigens isolated from a number of B. pseudomallei strains (Perry et al., 1995). Their results demonstrated that B. pseudomallei strains coordinately express two distinct somatic O-antigens (PS) on their cell surface. The type I antigen consists of a highmolecular-weight unbranched 1,3-linked homopolymer of 2-O-acetylated 6-deoxy-β-Dmanno-heptopyranosyl residues, while the type II antigen is an unbranched heteropolymer consisting of (-3)-β-D-glucopyranose-(1-3)-6deoxy-α-L-talopyranosyl-(1)-disaccharide repeats (L-6dTalp: __33% O-4 acetylated and O-2 methylated; __66% O-2 acetylated; Knirel et al., 1992; Perry et al., 1995). Even though the simultaneous expression of two or more LPS moieties is not an uncommon feature associated with Gram-negative bacteria, the degree to which the two PS antigens are conserved amongst B. pseudomallei strains is quite remarkable (Perry et al., 1995). It has been reported that B. pseudomallei strains are resistant to the bactericidal effects of normal human serum (Ismail et al., 1988). Type II PS is essential for conferring this resistance phenotype. A 17.5-kb region of the chromosome, which is required for the synthesis of the type II antigen, confers the serum-resistance phenotype (DeShazer et al., 1998). Type II PS is probably a significant determinant in the pathogenesis of melioidosis because the LD50 value associated with a type II PS mutant is approximately 140-

856

D.E. Woods and P.A. Sokol

fold higher than that of the wild-type strain (Woods et al., 1993; DeShazer et al., 1998). Flagella are commonly recognized as important virulence determinants expressed by bacterial pathogens because the motility phenotype imparted by these organelles often correlates with the ability of an organism to cause disease (Penn and Luke, 1992; Moens and Vanderleyden, 1996). A significant degree of size and antigenic homogeneity exists amongst flagellins expressed by B. pseudomallei isolates. Furthermore, flagellin-specific antiserum can passively protect diabetic infant rats against a B. pseudomallei challenge (Brett et al., 1994). Curiously, there was no significant difference between the virulence capacities associated with a wild-type strain of B. pseudomallei and nonmotile mutants in either the diabetic infant rat or Syrian hamster models of infection (DeShazer et al., 1997). These results indicate that though flagella and/or motility may not be major virulence determinants in the pathogenesis of melioidosis, purified flagellin might still be a protective immunogen against B. pseudomallei infections. Although glanders is a serious life-threatening zoonotic disease, relatively little is known about the pathogenesis, virulence factors, strain and the host immunopathological responses to infection (Fritz et al., 1999).

Treatment Burkholderia cepacia is resistant to most antibiotics, making treatment of CF patients very difficult. It usually is susceptible only to piperacillin, axlocillin, cefoperazone, ceftazidime, chloramphenicol and trimethoprim-sulfamethoxazole (TMP-SMX; Pitkin et al., 1997). Strains isolated from CF sputum from patients treated with several courses of antibiotic therapy are often resistant to all known antimicrobial agents (Gilligan and Whittier, 1999). Resistance to a variety of antimicrobial agents including penicillins, firstand second-generation cephalosporins and many of the aminoglycosides is characteristic of B. pseudomallei clinical isolates (Dance et al., 1988; Leelarasamee and Bovornkitti, 1989; Godfrey et al., 1991; Weinberg and Heller, 1997). With this in mind, accurate identification of the organism, evaluation of the severity of the infection and antibiotic susceptibility testing are of paramount importance in devising an effective chemotherapeutic strategy. Whereas the newer therapies that utilize combinations of ceftazidime–cotrimoxazole or amoxicillin–clavulanate for treatment of disease are proving beneficial, the mortality rates associated with the acute septicemic and pulmonary forms of melioidosis are still unacceptably high (Smith et al., 1987;

CHAPTER 3.2.16

Leelarasamee and Bovornkitti, 1989; Kanai and Kondo, 1994; Weinberg and Heller, 1997; Ho et al., 1997). Typically, prolonged oral therapy also is recommended to assure the full clinical resolution of infections while reducing the potential for recrudescence of disease. Antibiotic therapy for glanders is complex (Batmanov, 1991); however, there have been reports that ceftazidime, imipenem, doxycycline and ciprofloxacin are active against both B. mallei and B. pseudomallei (Kenny et al., 1999). Antibiotics clinically proven to be effective in the treatment of melioidosis may therefore be effective for treating glanders. Acknowledgements. This work was supported by Department of Defense Contract DAMD 17-98-C-8003, the Canadian Bacterial Diseases Network of Centres of Excellence, the Medical Research Council of Canada and the Canadian Cystic Fibrosis Foundation.

Literature Cited Ashdown, L. R., and J. M. Koehler. 1990. Production of hemolysin and other extracellular enzymes by clinical isolates of Pseudomonas pseudomallei. J. Clin. Microbiol. 28:2331–2334. Baird, R. M., H. Brown, A. W. Smith, and M. L. Watson. 1999. Burkholderia cepacia is resistant to the antimicrobial activity of airway epithelial cells. Immunopharmacology 44(3):267–272. Batmanov, V. P. 1991. Sensitivity of Pseudomonas mallei to fluoroquinolones and their efficacy in experimental glanders. Antibiot. Khimioter. 36:31–34. Bauernfeind, A., C. Roller, D. Meyer, R. Jungwirth, and I. Schneider. 1998. Molecular procedure for rapid detection of Burkholderia mallei and Burkholderia pseudomallei. J Clin Microbiol. 36(9):2737–2744. Bauernfeind, A., I. Schneider, R. Jungwirth, and C. Roller. 1999. Discrimination of Burkholderia multivorans and Burkholderia vietnamiensis from Burkholderia cepacia genomovars I, III and IV by PCR. J. Clin. Microbiol. 37:1335–1339. Benenson, A. S. 1995. Control of Communicable Diseases manual. American Public Health Association. Washington DC, 577. Bharadwaj, R., A. Kagal, S. K. Deshpandy, S. A. Joshi, P. M. Khare, A. R. Junnakar, and M. A. Phadke. 1994. Outbreak of plague-like illness caused by Pseudomonas pseudomallei in Maharashtra, India. Lancet 344:1574. Brett, P. J., D. C. Mah, and D. E. Woods. 1994. Isolation and characterization of Pseudomonas pseudomallei flagellin proteins. Infect. Immunol. 62:1914–1919. Brett, P. J., D. DeShazer, and D. E. Woods. 1997. Characterization of Burkholderia pseudomallei and Burkholderia pseudomallei-like strains. Epidemiol. Infect. 118:137– 148. Brett, P. J., D. DeShazer, and D. E. Woods. 1998. Burkholderia thailandensis sp. Nov., a Burkhoderia pseudomalleilike species. Int. J. System. Bacteriol. 48:317–320.

CHAPTER 3.2.16 Bryan, L. E., S. Wong, D. E. Woods, D. A. B. Dance, and W. Chaowagul. 1994. Passive protection of diabetic rats with antisera specific for the polysaccharide portion of the lipopolysaccharide isolated from Pseudomonas pseudomallei. Can. J. Infect. Dis. 5:170–178. Burkholder, W. H. 1950. Sour skin, a bacterial rot of onion bulbs. Phytopathology 40:115–117. Burns, J. L., M. Jonas, E. Y. Chi, D. K. Clark, A. Berger, and A. Griffith. 1996. Invasion of respiratory epithelial cells by Burkholderia (Pseudomonas) cepacia. Infect. Immunol. 64(10):4054–4059. Chaowagul, W., N. J. White, D. A. B. Dance, Y. Wattanagoon, P. Naigowit, T. M. E. Davis, S. Looareesuwan, and N. Pitawatchara. 1989. Melioidosis, a major cause of community-acquired septicemia in northeastern Thailand. J. Infect. Dis. 159:890–899. Corey, M., and V. Farewell. 1996. Determinants of mortality from cystic fibrosis in Canada, 1970–1989. Am. J. Epidemiol. 143(10):1007–1017. Crosa, J. H. 1989. Genetics and molecular biology of siderophore-mediated iron transport in bacteria. Microbiol. Rev. 53(4):517–530. Dance, D. A., V. Wuthiekanun, N. J. White, and W. Chaowagul. 1988. Antibiotic resistance in Pseudomonas pseudomallei. Lancet 1:994–995. Dance, D. A., V. Wuthiekanun, P. Naigowit, and N. J. White. 1989. Identification of Pseudomonas pseudomallei in clinical practice: use of simple screening tests and API 20NE. J. Clin. Pathol. 42:645–648. Dance, D. A. 1990. Melioidosis. Rev. Med. Microbiol. 1:143– 150. Dance, D. A. 1991. Melioidosis: The tip of the iceberg? Clin. Microbiol. Rev. 4:52–60. Darling, P., M. Chan, A. D. Cox, and P. A. Sokol. 1998. Siderophore production by cystic fibrosis isolates of Burkholderia cepacia. Infect. Immunol. 66(2):874–877. DeShazer, D., P. Brett, R. Carlyon, and D. E. Woods. 1997. Mutagenesis of Burkholderia pseudomallei with Tn 5OT182: isolation of motility mutants and molecular characterization of the flagellin structural gene. J. Bacteriol. 179:2116–2125. DeShazer, D., P. Brett, and D. E. Woods. 1998. The Type II O-antigen moiety of Burkholderia pseudomallei lipopolysaccharide is required for serum resistance and virulence. Molec. Microbiol. 30:1011–1081. Deshazer, D., and D. E. Woods. 1999. Pathogenesis of melioidosis: Use of Tn5-OT182 to study the molecular basis of Burkholderia pseudomallei virulence. J. Infect. Dis. Antimicrob. Ag. 16:91–96. Ellison, D. W., H. J. Baker, and M. Mariappan. 1969. Melioidosis in Malaysia. I: A method of isolation of Pseudomonas pseudomallei from soil and surface water. Am. J. Trop. Med. Hyg. 18:694–697. Esselman, M. T., and P. V. Liu. 1961. Lecithinase production by Gram-negative bacteria. J. Bacteriol. 81:939–945. Frangolias, D. D., E. Mahenthiralingam, S. Rae, J. M. Raboud, A. G. F. Davidson, R. Wittmann, and P. G. Wilcox. 1999. Burkholderia cepacia in cystic fibrosis: Variable disease course. Am. J. Respir. Crit. Care Med. 160:1572–1577. Freeman, B. A. 1985. Pseudomonas and Legionella. In: Burrow’s Textbook of Microbiology. W. B. Saunders Co. Philadelphia, PA. 544–557. Fritz, D. L, P. Vogel, D. R. Brown, and D. M. Waag. 1999. The hamster model of intraperitoneal Burkholderia mallei (glanders). Vet. Pathol. 36(4):276–291.

The Genus Burkholderia

857

Gilligan, P. H., P. A. Gage, L. M. Bradshaw, D. V. Schidlow, and B. T. DeCicco. 1985. Isolation medium for the recovery of Pseudomonas cepacia from respiratory secretions of patients with cystic fibrosis. J. Clin. Microbiol. 22:5–8. Gilligan, P. H. 1991. Microbiology of airway disease in patients with cystic fibrosis. Clin. Microbiol. Rev. 4(1):35–51. Gilligan, P. H., and S. Whittier. 1999. Burkholderia, Stenotrophomonas, Ralstonia, Brevundimonas, Comamonas, and Acidovorax. In: P. R. Murray, E. J. Baron, M. A. Pfaller, F. C. Tenover, and R. H. Yolken (Eds.) Manual of Clinical Microbiology. ASM Press. Washington DC. Gillis, M., T. V. Van, R. Bardin, M. Goor, P. Hebbar, A. Willems, P. Segers, K. Kersters, T. Heulin, and M. P. Fernandez. 1995. Polyphasic taxonomy in the genus Burkholderia leading to an amended description of the genus and proposition of Burkholderia vietnamiensis sp. nov. for N2-fixing isolates from rice in Vietnam. Int. J. Syst. Bacteriol. 45:274–289. Godfrey, A. J., S. Wong, D. A. Dance, W. Chaowagul, and L. E. Bryan. 1991. Pseudomonas pseudomallei resistance to beta-lactam antibiotics due to alterations in the chromosomally encoded beta-lactamase. Antimicrob. Agents Chemother. 35:1635–1640. Goldman, D. A., and J. D. Klinger. 1986. Pseudomonas cepacia: Biology, mechanisms of virulence, epidemiology. J. Pediatr. 108:806–812. Govan, J. R., P. H. Brown, J. Maddison, C. J. Doherty, J. W. Nelson, M. Dodd, A. P. Greening, and A. K. Webb. 1993. Evidence for transmission of Pseudomonas cepacia by social contact in cystic fibrosis. Lancet 342(8862):15–19. Govan, J. R., and V. Deretic. 1996a. Microbial pathogenesis in cystic fibrosis: Mucoid Pseudomonas aeruginosa and Burkholderia cepacia. Microbiol. Rev. 60(3):539–574. Govan, J. R., J. E. Hughes, and P. Vandamme. 1996b. Burkholderia cepacia: Medical, taxonomic and ecological issues. J. Med. Microbiol. 45(6):395–407. Haase, A., J. Janzen, S. Barrett, and B. Currie. 1997. Toxin production by Burkholderia pseudomallei strains and correlation with severity of melioidosis. J. Med. Microbiol. 46:557–563. Haubler, S., M. Nimtz, T. Domke, V. Wray, and I. Steinmetz. 1998. Purification and characterization of a cytotoxic exolipid of Burkholderia pseudomallei. Infect. Immunol. 66:1588–1593. Heckly, R. J., and C. Nigg. 1958. Toxins of Pseudomonas pseudomallei. II: Characterization. J. Bacteriol. 76:427– 436. Heckly, R. J. 1964. Differentiation of exotoxin and other biologically active substances in Pseudomonas pseudomallei filtrates. J. Bacteriol. 88:1730–1736. Henry, D. A., M. E. Campbell, J. J. LiPuma, and D. P. Speert. 1997. Identification of Burkholderia cepacia isolates from patients with cystic fibrosis and use of a simple new selective medium. J. Clin. Microbiol. 35:614–619. Henry, D. A., M. Campbell, C. McGimpsey, A. Clarke, L. Louden, J. Burns, M. H. Roe, P. Vandamme, and D. Speert. 1999. Comparison of isolation media for recovery of Burkholderia cepacia complex from respiratory secretions of patients with cystic fibrosis. J. Clin. Microbiol. 37:1004–1007. Ho, M., T. Schollaardt, M. D. Smith, M. B. Perry, P. J. Brett, W. Chaowagul, and L. E. Bryan. 1997. Specificity and functional activity of anti-Burkholderia pseudomallei polysaccharide antibodies. Infect. Immunol. 65:3648– 3653.

858

D.E. Woods and P.A. Sokol

Holmes, A., R. Nolan, R. Taylor, R. Finley, M. Riley, R. Z. Jiang, S. Steinbach, and R. Goldstein. 1999. An epidemic of Burkholderia cepacia transmitted between patients with and without cystic fibrosis. J. Infect. Dis. 179(5):1197–1205. Hornick, R. B. 1982. Diseases due to Pseudomonas mallei and Pseudomonas pseudomallei. In: R. J. Wedgewood (Ed.) Infections in Children. Harper & Row. Philadelphia, PA. 910–913. Howe, C. 1949. Glanders. In: H. A. Christian (Ed.) The Oxford Medicine. Oxford University Press. New York, NY. 185–201. Howe, C., A. Sampath, and M. Spotnitz. 1971. The Pseudomallei group: A review. J. Infect. Dis. 124:598– 606. Hutchison, M. L., I. R. Poxton, and J. R. Govan. 1998. Burkholderia cepacia produces a hemolysin that is capable of inducing apoptosis and degranulation of mammalian phagocytes. Infect. Immunol. 66(5):2033–2039. Inglis, T. J. J., S. C. Garrow, C. Adams, M. Henderson, and M. Mayo. 1998. Dry season outbreak of melioidosis in Western Australia. Lancet 352:1600. Isles, A., I. Maclusky, M. Corey, R. Gold, C. Prober, P. Fleming, and H. Levison. 1984. Pseudomonas cepacia infection in cystic fibrosis: An emerging problem. J. Pediatr. 104(2):206–210. Ismail, G., M. N. Embi, O. Omar, and N. Razak. 1987. Toxigenic properties of Pseudomonas pseudomallei extracellular products. Trop. Biomed. 4:101–110. Ismail, G., N. Razak, R. Mohamed, N. Embi, and O. Omar. 1988. Resistance of Pseudomonas pseudomallei to normal human serum bactericidal action. Microbiol. Immunol. 32:645–652. Johnson, W. M., S. D. Tyler, and K. R. Rozee. 1994. Linkage analysis of geographic and clinical clusters in Pseudomonas cepacia infections by multilocus enzyme electrophoresis and ribotyping. J. Clin. Microbiol. 32(4):924– 930. Kanai, K., and E. Kondo. 1994. Recent advances in biomedical sciences of Burkholderia pseudomallei (basonym: Pseudomonas pseudomallei). Jpn. J. Med. Sci. Biol. 47:1–45. Kenny, D. J., P. Russell, D. Rogers, S. M. Eley, and R. W. Titball. 1999. In vitro susceptibilities of Burkholderia mallei in comparison to those of other pathogenic Burkholderia spp.. Antimicrob. Agents Chemother. 43(11);Nov:2773–2775. Knirel, Y. A., N. A. Paramonov, A. S. Shashkov, N. K. Kochetkov, R. G. Yarullin, S. M. Farber, and V. I. Efremenko. 1992. Structure of the polysaccharide chains of Pseudomonas pseudomallei lipopolysaccharides. Carbohydr. Res. 233:185–193. Kooi, C., A. Cox, P. Darling, and P. A. Sokol. 1994. Neutralizing monoclonal antibodies to an extracellular Pseudomonas cepacia protease. Infect. Immunol. 62(7):2811–2817. Kovalev, G. K. 1971. Glanders (review). Zh. Mikrobiol. Edidemiol. Immunobiol. 48:63–70. Leelarasamee, A., and S. Bovornkitti. 1989. Melioidosis: Review and update. Rev. Infect. Dis. 11:413–425. Lewenza, S., B. Conway, E. P. Greenberg, and P. A. Sokol. 1999. Quorum sensing in Burkholderia cepacia: identification of the LuxRI homologs CepRI. J. Bacteriol. 181(3):748–756. LiPuma, J. J., J. E. Mortensen, S. E. Dasen, T. D. Edlind, D. V. Schidlow, J. L. Burns, and T. L. Stull. 1988.

CHAPTER 3.2.16 Ribotype analysis of Pseudomonas cepacia. J. Infect. Dis. 1164:133–136. LiPuma, J. J., S. E. Dasen, D. W. Neilson, R. C. Stern, and T. L. Stull. 1990. Person-to-person transmission of Pseudomonas cepacia between patients with cystic fibrosis. Lancet 336:527–532. LiPuma, J. J. 1998. Burkholderia cepacia: Management issues and new insights. Clin. Chest Med. 19(3):473–486, vi. Lonon, M. K., D. E. Woods, and D. C. Straus. 1988. Production of lipase by clinical isolates of Pseudomonas cepacia. J. Clin. Microbiol. 26(5):979–984. Mahenthiralingam, E., M. E. Campbell, D. A. Henry, and D. A. Speert. 1996. Epidemiology of Burkholderia cepacia infection in patients with cystic fibrosis: Analysis by random amplified polymorphic DNA fingerprinting. J. Clin. Microbiol. 34:2914–2920. Mahenthiralingam, E., D. A. Simpson, and D. P. Speert. 1997. Identification and characterization of a novel DNA marker associated with epidemic Burkholderia cepacia strains recovered from patients with cystic fibrosis. J. Clin. Microbiol. 35(4):808–816. Mahenthiralingam, E., T. Coenye, J. W. Chung, D. P. Speert, J. R. W. Govan, P. Taylor, and P. Vandamme. 2000. Diagnostically and experimentally useful panel of strains from the Burkholderia cepacia complex. J. Clin. Microbiol. 38(2):910–913. Martin, D. W., and C. D. Mohr. 2000. Invasion and intracellular survival of Burkholderia cepacia. Infect. Immunol. 68(1):24–29. Masoud, H., M. Ho, T. Schollaardt, and M. B. Perry. 1997. Characterization of the capsular polysaccharide of Burkholderia (Pseudomonas) pseudomallei 304b. J. Bacteriol. 179:5663–5669. Mays, E. E., and E. A. Ricketts. 1975. Melioidosis: Recrudescence associated with bronchogenic carcinoma twentysix years following initial geographic exposure. Chest 68:261–263. McCormick, J. B., D. J. Sexton, J. G. McMurray, E. Carey, P. Hayes, and R. A. Feldman. 1975. Human-to-human transmission of Pseudomonas pseudomallei. Ann. Internat. Med. 83:512–513. McKevitt, A. I., and D. E. Woods. 1984. Characterization of Pseudomonas cepacia isolates from patients with cystic fibrosis. J. Clin. Microbiol. 19(2):291–293. McKevitt, A. I., S. Bajaksouzian, J. D. Klinger, and D. E. Woods. 1989. Purification and characterization of an extracellular protease from Pseudomonas cepacia. Infect. Immunol. 57(3):771–778. Mietzner, T., and S. Morse. 1994. The role of iron-binding proteins in the survival of pathogenic bacteria. Ann. Rev. Nutr. 14:471–493. Moens, S. M., and J. Vanderleyden. 1996. Functions of bacterial flagella. Crit. Rev. Microbiol. 22:67–100. Mortensen, J. E., M. C. Fisher, and J. J. LiPuma. 1995. Recovery of Pseudomonas cepacia and other Pseudomonas species from the environment. Infect. Control. Hosp. Epidemiol. 16:30–32. Nakazawa, T., Y. Yamada, and M. Ishibashi. 1987. Characterization of hemolysin in extracellular products of Pseudomonas cepacia. J. Clin. Microbiol. 25(2):195– 198. Nigg, C., R. J. Heckly, and M. Colling. 1955. Toxin produced by Malleomyces pseudomallei. Proc. Soc. Exp. Biol. Med. 89:17–20. Nijhuis, E. H., M. J. Maat, I. W. E. Zeegers, C. Waalwijk, and J. A. Van-Venn, J.A. 1993. Selection of bacteria suitable

CHAPTER 3.2.16 for introduction into the rhizosphere of grass. Soil Biol. Biochem. 25:885–895. Nimtz, M., V. Wray, T. Domke, B. Brenneke, S. Haussler, and I. Steinmetz. 1997. Structure of an acidic exopolysaccharide of Burkholderia pseudomallei. Eur. J. Biochem. 250:608–616. Palleroni, N. J., and B. Holmes. 1981. Pseudomonas cepacia sp. nov., nom. rev. Int. J. Syst. Bacteriol. 31:479–481. Parker, M. T. 1990. Glanders and melioidosis. In: M. T. Parker and L. H. Collier (Eds.) Topley & Wilson’s Principles of Bacteriology, Virology and Immunity. B. C. Decker Inc. Philadelphia, PA. 392–394. Pegues, D. A., L. A. Carson, O. C. Tablan, S. C. FitzSimmons, S. B. Roman, J. M. Miller, and W. R. Jarvis. 1994. Acquisition of Pseudomonas cepacia at summer camps for patients with cystic fibrosis: Summer Camp Study Group. J. Pediatr. 124(5, Part 1):694–702. Pegues, C. F., D. F. Pegues, D. S. Ford, P. L. Hibberd, L. A. Carson, C. M. Raine, and D. C. Hooper. 1996. Burkholderia cepacia respiratory acquisition: epidemiology and molecular characterization of a large nosocomial outbreak. Epidemiol. Infect. 116:309–317. Penn, C. W., and C. J. Luke. 1992. Bacterial flagellar diversity and significance in pathogenesis. FEMS Microbiol. Lett. 100:331–336. Perry, M. B., L. L. MacLean, T. Schollaardt, L. E. Bryan, and M. Ho. 1995. Structural characterization of the lipopolysaccharide O antigens of Burkholderia pseudomallei. Infect. Immunol. 63:3348–3352. Pitkin, D. H., W. Sheikh, and H. L. Nadler. 1997. Comparative in vitro activity of meropenem versus other extended-spectrum antimicrobials against randomly chosen and selected resistant clinical isolates tested in 26 North American centers. Clin. Infect. Dis. 24(Suppl. 2):S238–S248. Pitt, T. L. 1990. Pseudomonas mallei and P. pseudomallei. In: M. T. Parker and L. H. Collier (Eds.) Topley & Wilson’s Principles of Bacteriology, Virology and Immunity. B. C. Decker Inc. Philadelphia, PA. 265–268. Pitt, T. L., H. Aucken, and D. A. Dance. 1992. Homogeneity of lipopolysaccharide antigens in Pseudomonas pseudomallei. J. Infect. 25:139–146. Prince, A. 1986. Antibiotic resistance of Pseudomonas species. J. Pediatr. 108:830–834. Pruksachartvuthi, S., N. Aswapokee, and K. Thakerngpol. 1990. Survival of Pseudomonas pseudomallei in human phagocytes. J. Med. Microbiol. 31:109–114. Pugsley, A. P. 1993. The complete general secretory pathway in Gram-negative bacteria. Microbiol. Rev. 57:50–108. Ralston, E., N. J. Palleroni, and M. Doudoroff. 1973. Pseudomonas pickettii, a new species of clinical origin related to Pseudomonas solanacearum. Int. J. Syst. Bacteriol. 23:15–19. Riboli, A. C., R. Koshinski, K. Arias, K. Marks-Austin, D. Stiertz, and T. L. Stull. 1996. An outbreak of Burkholderia cepacia lower respiratory tract infection associated with contaminated albuterol nebulization solution. Infect. Control Hosp. Epidemiol. 17:741–743. Saini, L. S., S. B. Galsworthy, M. A. John, and M. A. Valvano. 1999. Intracellular survival of Burkholderia cepacia complex isolates in the presence of macrophage cell activation. Microbiology 145(12):3465–3475. Sanford, J. P. 1990. Principles and Practice of Infectious Diseases. In: G. L. Mandell, R. G. Douglas Jr., and J. E. Bennett (Eds.) Churchill Livingstone. New York, NY. 1692–1696.

The Genus Burkholderia

859

Sanford, J. P. 1995. Pseudomonas species including melioidosis and glanders. In: G. L. Mandell, J. E. Bennett and R. Dolin (Eds.) Principles and Practice of Infectious Diseases, 4th ed. Churchill Livingstone. New York, NY. 1250–1254. Segonds, C., T. Heulin, N. Marty, and G. Chabanon. 1999. Differentiation of Burkholderia species by PCRrestriction fragment length polymorphism analysis of the 16S rRNA gene and application to cystic fibrosis isolates. J. Clin. Microbiol. 37:2201–2208. Sexton, M. M., A. L. Jones, W. Chaowagul, and D. E. Woods. 1994. Purification and characterization of a protease from Pseudomonas pseudomallei. Can. J. Microbiol. 40:903–910. Simpson, I. N., J. Finlay, D. J. Winstanley, N. Dewhurst, J. W. Nelson, S. L. Butler, and J. R. Govan. 1994. Multiresistance isolates possessing characteristics of both Burkholderia (Pseudomonas) cepacia and Burkholderia gladioli from patients with cystic fibrosis. J. Antimicrob. Chemother. 34(3):353–361. Smith, C. J., J. C. Allen, M. N. Embi, O. Othman, N. Razak, and G. Ismail. 1987. Human melioidosis: An emerging medical problem. MIRCEN J. 3:343–366. Smith, D. L., L. B. Gumery, E. G. Smith, D. E. Stableforth, M. E. Kaufmann, and T. L. Pitt. 1993. Epidemic of Pseudomonas cepacia in an adult cystic fibrosis unit: Evidence of person-to-person transmission. J. Clin. Microbiol. 31(11):3017–3022. Smith, M. D., V. Wuthiekanun, A. L. Walsh, and N. J. White. 1995. Quantitative recovery of Burkholderia pseudomallei from soil in Thailand. Trans. R. Soc. Trop. Med. Hyg. 89:488–490. Sokol, P. A. 1986. Production and utilization of pyochelin by clinical isolates of Pseudomonas cepacia. J. Clin. Microbiol. 23(3):560–562. Sokol, P. A., P. Darling, D. E. Woods, E. Mahenthiralingam, and C. Kooi. 1999. Role of ornibactin biosynthesis in the virulence of Burkholderia cepacia: characterization of pvdA, the gene encoding L-ornithine N(5)-oxygenase. Infect. Immunol. 67(9):4443–4455. Sokol, P. A., C. Kooi, R. S. Hodges, P. Cachia, and D. E. Woods. 2000. Immunization with a Pseudomonas elastase peptide reduces severity of experimental lung infections due to Pseudomonas aeruginosa or Burkholderia cepacia. J. Infect. Dis. 181:1682–1692. Speert, D. P., M. Bond, R. C. Woodman, and J. T. Curnutte. 1994. Infection with Pseudomonas cepacia in chronic granulomatous disease: Role of nonoxidative killing by neutrophils in host defense. J. Infect. Dis. 170(6):1524– 1531. Steele, J. H. 1973. The zoonoses: An epidemiologist’s viewpoint. In: M. Stefanini (Ed.) Progress in Clinical Pathology. Grune and Stratton. New York, NY. 239– 286. Steinmetz, I., M. Rohde, and B. Brenneke. 1995. Purification and characterization of an exopolysaccharide of Burkholderia (Pseudomonas) pseudomallei. Infect. Immunol. 63:3959–3965. Sun, L., R. Z. Jiang, S. Steinbach, A. Holmes, C. Campanelli, J. Forstner, U. Sajjan, Y. Tan, M. Riley, and R. Goldstein. 1995. The emergence of a highly transmissible lineage of cbl+ Pseudomonas (Burkholderia) cepacia causing CF centre epidemics in North America and Britain. Nat. Med. 1(7):661–666. Tanphaichitra, D., and S. Srimuang. 1984. Cellular immunity in tuberculosis, melioidoses, pasteurellosis, penicilliosis

860

D.E. Woods and P.A. Sokol

and role of levamisole and isoprinosine. Dev. Biol. Stand. 57:117–123. Tanphaichitra, D. 1989. Tropical disease in the immunocompromised host: melioidosis and pythiosis. Rev. Infect. Dis. 11(Suppl. 7):S1629–S1643. Vandamme, P., B. Holmes, M. Vancanneyt, T. Coenye, B. Hoste, R. Coopman, H. Revets, S. Lauwers, M. Gillis, K. Kersters, and J. R. Govan. 1997. Occurrence of multiple genomovars of Burkholderia cepacia in cystic fibrosis patients and proposal of Burkholderia multivorans sp. nov. Int. J. Syst. Bacteriol. 47(4):1188–1200. Vandamme, P., E. Mahenthiralingam, B. Holmes, T. Coenye, B. Hoste, P. De Vos, D. Henry, and D. P. Speert. 2000. Identification and population structure of Burkholderia stabilis sp. nov (formerly Burkholderia cepacia genomovar IV). J. Clin. Microbiol. 38:1042–1047. Von Graevenitz, A. 1973. Clinical microbiology of unusual Pseudomonas species. Prog. Clin. Path. 5: 185–218. Weinberg, A. N., and H. M. Heller. 1997. Infectious Diseases of the Lung. Thieme-Stratton. New York, NY. 2413– 2430. Welch, D. F., M. J. Muszynski, C. H. Pai, M. J. Maracon, M. M. Hribar, P. H. Gilligan, J. M. Matsen, P. A. Ahlin, B. C. Hilman, and S. A. Chartrand. 1987. Selective and differential medium for recovery of Pseudomonas cepacia from the respiratory tracts of patients with cystic fibrosis. J. Clin. Microbiol. 25:1730–1734. Whitmore, A., and C. S. Krishnaswami. 1912. An account of the discovery of a hitherto undescribed infective disease occurring among the population of Rangoon. Indian Med. Gaz. 47:262–267. Whitmore, A. 1913. An account of a glanders-like disease occurring in Rangoon. J. Hyg. Camb. 13:1–7.

CHAPTER 3.2.16 Woods, D. E., A. L. Jones, and P. J. Hill. 1993. Interaction of insulin with Pseudomonas pseudomallei. Infect. Immunol. 61:1914–1919. Wuthiekanum, V., M. D. Smith, D. A. B. Dance, A. L. Walsh, T. L. Pitt, and N. J. White. 1996a. Biochemical characteristics of clinical and environmental isolates of Burkholderia pseudomallei. J. Med. Microbiol. 45:408– 412. Wuthiekanum, V., M. D. Smith, D. A. B. Dance, A. L. Walsh, T. L. Pitt, and N. J. White. 1996b. Isolation of Pseudomonas pseudomallei from soil in north-eastern Thailand. Trans. R. Soc. Trop. Med. Hyg. 89:41–43. Yabuuchi, E., Y. Kosako, H. Oyaizu, I. Yano, H. Hotta, Y. Hashimoto, T. Ezaki, and M. Arakawa. 1992. Proposal of Burkholderia gen. nov. and transfer of seven species of the genus Pseudomonas homology group II to the new genus, with the type species Burkholderia cepacia (Palleroni and Holmes 1981) comb. nov. Microbiol. Immunol. 36(12):1251–1275. Yabuuchi, E., Y. Kosako, I. Yano, H. Hotta, and Y. Nishiuchi. 1995. Transfer of two Burkholderia and an Alcaligenes species to Ralstonia gen. nov.: Proposal of Ralstonia pickettii (Ralston, Palleroni and Doudoroff 1973) comb. nov., Ralstonia solanacearum (Smith 1896) comb. nov. and Ralstonia eutropha (Davis 1969) comb. nov. Microbiol. Immunol. 39:897–904. Yang, H. M., W. Chaowagul, and P. A. Sokol. 1991. Siderophore production by Pseudomonas pseudomallei. Infect. Immunol. 59:776–780. Yang, H. M., C. D. Kooi, and P. A. Sokol. 1993. Ability of Pseudomonas pseudomallei malleobactin to acquire transferrin-bound, lactoferrin-bound, and cell-derived iron. Infect. Immunol. 61:656–662.

Prokaryotes (2006) 5:861–872 DOI: 10.1007/0-387-30745-1_41

CHAPTER 3.2.17 ehT

e t i r t iN

gn i z i d i xO

a i re t caB

The Nitrite-Oxidizing Bacteria AHARON ABELIOVICH

Introduction Nitrite-oxidizing bacteria are a small group of primarily organo and/or chemoautotrophs that are difficult to grow and work with. For many years they drew very little attention, although it was realized that they provide a key link in the global nitrogen cycle between the ammoniaoxidizing bacteria, which generate nitrite, and the various denitrifying microorganisms that remove nitrate by reducing it to ammonia or molecular nitrogen, thus completing the global nitrogen cycle. The growing public awareness relating to issues of environmental pollution in recent decades brought renewed interest in this group of bacteria, as their role in the transformation of nitric oxides became apparent. In addition, these bacteria cause economic damage to agriculture and to ground water quality by contributing to the leaching of nitrogen fertilizers (distributed in the form of ammonia) from surface soils, thus polluting aquifers with increasing concentrations of nitrites and nitrates. Owing to their key role in the global cycling of nitrogen and their activities both as prime contributors to and scavengers of nitric oxides in the biosphere, nitrite-oxidizing bacteria became in recent years the focus of intensive ecophysiological research, in spite of the difficulties in cultivating and maintaining many of these bacteria in the laboratory. In addition to this group of autotrophic nitrifyers, there is also a diverse group of heterotrophic bacteria capable of heterotrophic nitrite oxidation. Thus, in one such study (Sakai et al., 1996), the nitrite transforming activities of heterotrophic bacterial strains from various culture collections as well as isolates from activated sludge were studied. Of the 48 strains tested, 17 strains consumed 1–5 mM of nitrite and accumulated corresponding amounts of nitrate. Heterotrophic microorganisms are rather flexible in determining the fate of the consumed nitrite. For instance, in Bacillus subtilis strain I-41 (a denitrification-positive isolate), the ratio of the amount

of nitrate accumulated to that of nitrite consumed varied from 0 to 100%, depending on the culture conditions. However, the mechanism of oxidation of nitrite to nitrate in these heterotrophic bacteria is very different from that in the autotrophic bacteria (Sorokin, 1991), as heterotrophic nitrite oxidation requires catalase and hydrogen peroxide (H2O2) generated through oxidation of organic electron donors. Also, aeration affects the nitrite- and nitrate-transforming activities of various heterotrophic bacteria. For example, Pseudomonas pavonaceae, a denitrification-positive strain, metabolizes both nitrite and nitrate to more reduced compounds at low oxygen pressure, and the direction of the conversion changes from reduction to oxidation at high oxygen pressure. This switching might be caused by inhibition and repression of the nitrite-reducing activity and by stimulation of nitrite-oxidizing activity by oxygen (Sakai et al., 1997).

Historic Background Nitrite-oxidizing bacteria carry the second stage of the nitrification process, that of oxidation of nitrite to nitrate. Because nitrate is an essential ingredient of gunpowder, nitrate and nitrification were the focus of interest both to scientists and politicians for many centuries. Nitrification protocols for the manufacture of nitrates by composting organic matter in soil have existed for many centuries: from the tenth century in China and from the twelfth century in Europe (Macdonald, 1986). However, the process was considered to be of a chemical and not biological nature until the late nineteenth century. It was only in 1862 that Pasteur suggested that nitrification was of biological origin (Pasteur, 1862), and it was only when Winogradsky in 1891 succeeded in isolating a nitrite-oxidizing bacterium that the debate about whether nitrification is a one-stage process (carried by a single bacterium) or a twostage process (carried by two distinct classes of microorganisms) was settled (Macdonald, 1986).

862

A. Abeliovich

CHAPTER 3.2.17

Nitrite Oxidation A detailed account of the mechanism of nitrite oxidation as an energy source has been described previously (Bock et al., 2001). The key enzyme that makes the nitrite-oxidizing bacteria so important in the global nitrogen cycle is nitrite oxidoreductase (NOR), which carries out the stoichiometric reaction: -

-

NO 2 + H 2O ¤ NO3 + 2H + + e -

(1)

2H + + 2e - + 0.5O 2 Æ H 2O

(2)

Reaction (1) is reversible, and many nitrite oxidizers are capable of reducing nitrate to nitrite under anaerobic conditions, but the significance of the reverse reaction to their survival is not clear, as both nitrite and nitrate are rapidly reduced in anaerobic environments. Nitrite oxidoreductase was isolated from various Nitrobacter strains, such as from mixotrophically grown cells of Nitrobacter hamburgensis; the enzyme purified from heat-treated membranes was homogeneous by the criteria of polyacrylamide gel electrophoresis and size exclusion chromatography. The monomeric form consisted of two subunits with molecular weights of 115 kDa and 65 kDa, respectively. The dimeric form of the enzyme contained 0.70 molybdenum, 23.0 iron, 1.76 zinc and 0.89 copper gram-atoms per molecule. The catalytically active enzyme was investigated by visible and electron paramagnetic resonance spectroscopy (EPR) under oxidizing (as isolated), reducing (dithionite), and turnover (nitrite) conditions. As isolated, the enzyme exhibited a complex set of EPR signals between 5–75 K, originating from several ironsulfur and molybdenum (V) centers. Addition of the substrate nitrite or the reducing agent dithionite resulted in a set of new resonances. The molybdenum and the iron-sulfur centers of nitrite oxidoreductase from Nitrobacter hamburgensis were involved in the transformation of nitrite to nitrate (Meincke et al., 1992).

Systematics As late as 1981, all bacteria capable of chemolithotrophic growth using ammonia ornitrite as energy source and capable of assimilating CO2 via the Calvin cycle were grouped in one family, the Nitrobacteraceae (Watson et al., 1981). This classification was based on the fact that all the bacteria grouped in this family carry out oxidation reactions of inorganic nitrogen compounds, and all are capable of chemolithotrophic growth using these as energy sources. All are Gramnegative bacteria and usually found in associ-

ation; as a physiological interdependence exists between the two groups, inasmuch as the ammonia oxidizers provide the substrate for the nitrite oxidizers, whereas the nitrite oxidizers are inhibited by excess ammonia. With the advent of molecular biology and classification of bacteria through sequencing of 16S ribosomal RNA genes and other specific DNA sequences, as well as the use of monoclonal antibodies, it became evident that the two groups are not related, and that there is no phylogenetic relationship between ammonia and nitrite oxidizers. Almost all nitrite oxidizers (with the exception of Nitrospira moscoviensis, which seems to occupy an intermediate position; Schramm et al., 1998) are a homogenous group belonging to the α-subdivision of the Proteobacteria and are very closely related to the nonsulfur purple photosynthetic bacterium Rhodopseudomonas palustris. None of the ammonia-oxidizing bacteria belong to the αsubdivision, and they are all distributed between the β- and γ-subdivisions (Woese et al., 1984; Woese et al., 1985). Comparative 16S rDNA sequencing was used to evaluate phylogenetic relationships among selected strains of ammonia- and nitriteoxidizing bacteria (Teske et al., 1994). All characterized strains were shown to be affiliated with the proteobacteria. The study extended recent 16S rDNA-based studies of phylogenetic diversity among nitrifiers by the comparison of eight strains of the genus Nitrobacter and representatives of the genera Nitrospira and Nitrospina. The latter genera were shown to be affiliated with the δ-subdivision of the proteobacteria but did not share a specific relationship to each other or to other members of the δ-subdivision. All characterized Nitrobacter strains constituted a closely related assemblage within the αsubdivision of the Proteobacteria. As previously observed, all ammonia-oxidizing genera except Nitrosococcus oceanus constitute a monophyletic assemblage within the β-subdivision of the proteobacteria. Consideration of physiology and phylogenetic distribution suggests that nitriteoxidizing bacteria of the α- and γ-subdivisions are derived from immediate photosynthetic ancestry. Each nitrifier retains the general structural features of the specific ancestor’s photosynthetic membrane complex. Thus, the nitrifiers, as a group, apparently are not derived from an ancestral nitrifying phenotype. Watson et al. (1981) identified three genera of nitrite-oxidizing bacteria (the Nitrospira being discovered only in 1986; Watson, 1986) and classified them on the basis of morphology and nutritional requirements. Bartosch et al. (1999) also divided the nitrite-oxidizing bacteria into four genera, Nitrobacter, Nitrospina, Nitrococcus and

CHAPTER 3.2.17

Nitrospira, according to their reactions with monoclonal antibodies prepared against the nitrite oxidoreductase of the genus Nitrobacter. Among these, Nitrobacter and Nitrospira strains are capable of heterotrophic or mixotrophic growth, the rest being strict chemolithotrophs. Because of the difficulties in culturing nitrifying bacteria on solid media, the methods for enumerating and identifying nitrifiers in environmental samples conventionally depended on incubating serial dilutions for very long periods, and then testing these for appearance of nitrate. Thus, the dominant nitrite-oxidizing species in an environmental sample was usually identified only after incubation of 3–6 months and was acertained on the basis of morphological and physiological properties such as sensitivity to salts and nutrient concentration. These observations led over the years to a general consensus that Nitrobacter strain were the dominant nitriteoxidizing organisms in practically all tested environments such as soil, marine waters, wastewater treatment plants, fresh water and reservoirs; Nitrococcus and Nitrospina were rarely identified (Watson et al., 1981). In 1995, a Gram-negative, nonmotile, nonmarine, nitrite-oxidizing bacterium was isolated by enrichment culture of a sample from a partially corroded area of an iron pipe located in a heating system in Moscow, Russia (Ehrich et al., 1995). The cells (0.9–2.2 µm × 0.2–0.4 µm in size) were helical to vibroid-shaped and often formed spirals with up to three turns 0.8–1.0 µm in width. The organism possessed an enlarged periplasmic space and lacked intracytoplasmic membranes and carboxysomes. The cells excreted extracellular polymers and formed aggregates. The bacterium grew optimally at 39°C and pH 7.6–8.0 in a mineral medium with nitrite as sole energy source and carbon dioxide as sole carbon source. The doubling time was 12 h in a mineral medium with 7.5 mM nitrite. The cell yield was low; only 0.9 mg of protein/liter was formed during oxidation of 7.5 mM nitrite. Under anoxic conditions, hydrogen could be used as electron donor with nitrate as electron acceptor. Organic matter (yeast extract, meat extract and peptone) supported neither mixotrophic nor heterotrophic growth. At concentrations as low as 0.75 g of organic matter/liter or higher, growth of the nitrite-oxidizing cells was inhibited. The cells contained cytochromes of the b- and c-type. The G+C content of DNA was 56.9 ± 0.4 mol%. This chemolithoautotrophic nitrite-oxidizer differed from the terrestrial members of the genus Nitrobacter with regard to morphology and substrate range and was similar to Nitrospira marina in both characteristics. The isolated bacterium was designated as a new species of the genus Nitrospira. Recent studies suggest that this genus

The Nitrite-Oxidizing Bacteria

863

dominates most of the nitrite oxidation activities in natural as well as artificial habitats (see below). With the introduction of molecular technologies to environmental microbiology, the true dominant nitrite oxidizers could be identified in situ in environmental samples, either by polymerase chain reaction (PCR, using specific rDNA primers) or fluorescence in situ hybridization (FISH, using specific probes). These studies led to the surprising conclusion that Nitrospira strains are the dominant nitrite oxidizers in most environmental samples tested so far, whereas Nitrobacter sp. are seldom identified without prior enrichment (Burrell et al., 1998; Burrell et al., 1999; Schramm et al., 1998; Juretschko et al.,1998; Okabe et al., 1999). There is, however, no doubt that within the next years, with the advent of oligonucleotide microchip technology, oligonucleotide microchips will dominate as genosensors for determinative and environmental studies in microbiology (Guschin et al., 1997). Precise identification of bacterial communities and of the genes switched on by each member of these communities under various environmental conditions will become routine in the environmental microbiology lab, and these procedures will enable the fine resolution of the community of nitrifying bacteria and its activities in different biotopes. DNA sequencing methodologies also were used to identify bacteria in situ, in biofilms and activated sludge particles (Schramm et al., 1998). Bacterial aggregates from a chemolithoautotrophic, nitrifying fluidized bed reactor were investigated with microsensors and rDNA-based molecular techniques. The microprofiles of O2, NH4+, NO2− and NO3− demonstrated the occurrence of complete nitrification in the outer 125 µm of the aggregates. FISH analysis showed that the dominant populations were of Nitrospira spp. and Nitrosospira sp. and that they formed separate, dense clusters which were in contact with each other and occurred throughout the aggregate. Significantly, no ammonia- or nitriteoxidizing bacteria of the genera Nitrosomonas or Nitrobacter, respectively, could be detected by FISH. To identify the nitrite oxidizers, a 16S ribosomal DNA clone library was constructed and screened by denaturing gradient gel electrophoresis (DGGE), and selected clones were sequenced. The organisms represented by these sequences formed two phylogenetically distinct clusters affiliated with the nitrite oxidizer Nitrospira moscoviensis. The dominant Nitrospira sp. formed clusters with the nitrosospira sp. and occurred throughout the aggregate while the second, smaller, morphologically and genetically different population of Nitrospira sp. was

864

A. Abeliovich

restricted to the outer nitrifying zones. The phylogeny of bacteria belonging to the genus Nitrobacter was investigated by sequencing the whole 16S rRNA gene (Orso et al., 1994). The average level of similarity for three Nitrobacter strains examined was high (99.2%), and the similarity level between Nitrobacter winogradsky and Nitrobacter sp. strain LL, which represent two different genomic species, was even higher (99.6%). When all of the Nitrobacter strains and their phylogenetic neighbors Bradyrhizobium and Rhodopseudomonas species were considered, the average similarity level was 98.1%. When complete sequences were used, Nitrobacter hamburgensis clustered with the two other Nitrobacter strains, though this was not the case when partial sequences were used. The two Rhodopseudomonas palustris strains examined exhibited a low similarity level (97.6%) and were not clustered. In another study (Okabe et al., 1999), the in situ spatial organization of ammonia-oxidizing and nitrite-oxidizing bacteria in domestic wastewater biofilms and autotrophic nitrifying biofilms was investigated by using microsensors and FISH, performed with 16S rDNA-targeted oligonucleotide probes. The combination of these techniques made it possible to relate in situ microbial activity directly to the occurrence of specific nitrifying bacterial populations. In situ hybridization revealed that bacteria belonging to the genus Nitrobacter were not detected; instead, Nitrospira-like bacteria were the main nitriteoxidizing bacteria in both types of biofilms. Nitrospira-like cells formed irregularly shaped aggregates consisting of small microcolonies, which bound the clusters of ammonia oxidizers. Whereas most of the ammonia-oxidizing bacteria were present throughout the biofilms, the nitrite-oxidizing bacteria were restricted to the active nitrite-oxidizing zones, which were inside the biofilms. Microelectrode measurements showed that the active ammonia-oxidizing zone was located in the outer part of a biofilm, whereas the active nitrite-oxidizing zone was located just below the ammonia-oxidizing zone and overlapped the location of nitrite-oxidizing bacteria, as determined by FISH. The use of fluorescent monoclonal antibodies was also very useful for the rapid quantification and in situ detection of specific nitrifiers in a mixed bacterial habitat such as a biofilm (Noda et al., 2000). In that study, 12 monoclonal antibodies against Nitrosomonas europaea (IFO14298) and 16 against Nitrobacter winogradsky (IFO 14297) enabled a direct cell count of N. europaea and N. winogradsky. Moreover, the distribution of N. europaea and N. winogradsky in a biofilm could be examined. Most of N. winogradsky existed near the surface and most

CHAPTER 3.2.17

of N. europaea existed within the polyethylene glycol (PEG) gel pellet. Using DNA sequences from the intergenic spacer (IGS) region of the ribosomal operon, and two primers derived from 16S and 23S rDNA conserved sequences, Navarro et al. (1992a) amplified these sequences by PCR. The PCR products, cleaved by four base cutting restriction enzymes, were used to differentiate Nitrobacter strains. This method is convenient for the genotypic characterization of Nitrobacter isolates and was successfully used to characterize natural populations of Nitrobacter from various soils and a lake. Considerable diversity was demonstrated in various soils, and in both water and sediments of the lake. Navarro et al. (1992b) also studied the genomic diversity of Nitrobacter strains by determining rDNA gene restriction patterns as well as hybridization characteristics and DNA base compositions. The DNA hybridization (S1 nuclease method) revealed five DNA genomic groups, and these groups formed three genomic Nitrobacter species. Mobarry et al. (1996) prepared a hierarchical set of five 16S rRNA-targeted DNA probes for phylogenetically defined groups of autotrophic ammonia- and nitrite-oxidizing bacteria. Their environmental application was demonstrated by quantitative slot blot hybridization and wholecell hybridization of nitrifying activated sludge and biofilm samples. In situ hybridization experiments revealed that Nitrobacter and Nitrosomonas species occurred in clusters and frequently were in contact with each other within sludge flocs. The nitrite oxidoreductase (NOR) from the facultative nitrite-oxidizing bacterium Nitrobacter hamburgensis X14 was used to develop a probe for the gene norB (Kirstein and Bock, 1993). Sequence analysis of DNA fragments revealed three adjacent open reading frames in the order norA, norX and norB. The deduced amino acid sequence of protein NorB contained four cysteine clusters with striking homology to those of iron-sulfur centers of bacterial ferredoxins. Protein NorB shares significant sequence similarity to the β-subunits (NarH and NarY) of the two dissimilatory nitrate reductases (NRA and NRZ) of Escherichia coli. Additionally, the derived amino acid sequence of the truncated open reading frame of norA showed striking resemblance to the α-subunits (NarG and NarZ) of the E. coli nitrate reductases. Additionally, the derived amino acid sequence of the truncated open reading frame of norA showed resemblance to the α-subunits of the E. coli nitrate reductases. Monoclonal antibodies prepared against nitrite oxidoreductases of nitrite oxidizers also

CHAPTER 3.2.17

were used successfully for the identification of nitrite oxidizers (Bartosch et al., 1999). Immunoblot analyses performed with three monoclonal antibodies (MAbs) that recognized the nitrite oxidoreductase (NOR) of the genus Nitrobacter were used for taxonomic investigations of nitrite oxidizers. These MAbs were able to detect the nitrite-oxidizing systems (NOS) of the genera Nitrospira, Nitrococcus and Nitrospina. When the genus-specific reactions of the MAbs were correlated with 16S rDNA sequences, they reflected the phylogenetic relationships among the nitrite oxidizers. In ecological studies, the immunoblot analyses demonstrated that Nitrobacter or Nitrospira cells could be enriched from activated sludge by using various substrate concentrations. The microbiology of the biomass from a nitrite-oxidizing sequencing batch reactor (NOSBR) as well as the seed sludge used were investigated by microscopy, by culturedependent methods, and by molecular biological methods (Burrell et al., 1998). The NOSBR was fed with an inorganic salts solution and nitrite as the sole energy source. It was operated for 6 months, and 16S ribosomal DNA clone libraries were prepared both from the seed sludge and from the reactor. Analysis of the seed sludge revealed that it contained three clones (4% of biomass) that were closely related to the autotrophic nitrite-oxidizer Nitrospira moscoviensis, whereas the NOSBR sludge itself was overwhelmingly dominated by bacteria closely related to the N. moscoviensis (89%). Only two clone sequences were similar to those of the genus Nitrobacter. Near-complete insert sequences of eight clones of N. moscoviensis isolated from the NOSBR and one clone from the seed sludge were determined and phylogenetically analyzed. This report was the first to show the presence of bacteria from the Nitrospira genus in wastewater treatment systems. Nitrite-oxidizing bacteria belonging exclusively to the genus Nitrospira also dominated the nitrite-oxidizing community of a phosphateremoving biofilm from a sequencing batch biofilm reactor (Gieseke et al., 2001). In another study (Burrell et al., 1999), a sequencing batch reactor (SBR) was operated to selectively grow a nitrite-oxidizing microbial community, and it was found that the nitrite oxidation was due the presence of bacteria from the Nitrospira genus and not the Nitrobacter genus, which were in very low numbers. It was hypothesized that the unknown nitrite-oxidizing bacteria in wastewater treatment plants are a range of species related to Nitrospira moscoviensis. Oxidation of nitrite to nitrate in aquaria is typically attributed to bacteria belonging to the genus Nitrobacter that are members of the α-

The Nitrite-Oxidizing Bacteria

865

subdivision of the class Proteobacteria. To identify bacteria responsible for nitrite oxidation in aquaria, clone libraries of rRNA genes were developed by Hovanec et al. (1998) from biofilms from several freshwater aquaria, and analysis of the rDNA libraries, along with results from denaturing gradient gel electrophoresis (DGGE) on frequently sampled biofilms, indicated the presence of putative nitrite-oxidizing bacteria closely related to other members of the genus Nitrospira. Hybridization experiments with rRNA from biofilms of freshwater aquaria demonstrated that Nitrospira-like rRNA comprised nearly 5% of the rRNA extracted from the biofilms during the establishment of nitrification. Nitrite-oxidizing bacteria belonging to the α-subdivision of the class Proteobacteria (e.g., Nitrobacter spp.) were not detected in these samples. Aquaria that received a commercial preparation containing Nitrobacter species did not show evidence of Nitrobacter growth and development but did develop substantial populations of Nitrospira-like species. Time series analysis of rDNA phylotypes on aquaria biofilms by DGGE, combined with nitrite and nitrate analysis, showed a correspondence between the appearance of Nitrospira-like bacterial ribosomal DNA and the initiation of nitrite oxidation. In total, the data suggest that Nitrobacter winogradsky and close relatives were not the dominant nitrite-oxidizing bacteria in freshwater aquaria. Instead, nitrite oxidation in freshwater aquaria appeared to be mediated by bacteria closely related to Nitrospira sp.

Isolation and Nutritional Requirements The information in this section is compiled from the chapter by Bock and Koops (1992) on Nitrobacter in the second edition published in the previous edition of {The Prokaryotes}. Obligate and facultative lithoautotrophic nitrite oxidizers can be successfully isolated only if mineral nitrite media free of any organic contaminants are used, otherwise heterotrophs will overgrow nitrite oxidizers. A sample of 1–2 g (soil, stone, mud etc.) is placed in 100 ml of enrichment medium selective for the growth of either terrestrial or marine strains (Table 1). For the enrichment of dominant nitrifiers, a combination of the most probable number (MPN) technique followed by the serial dilutions method is suitable. In the highest dilutions, nitrite oxidation can be detected only after several months because of the slow growth of the nitrite oxidizers. Plating samples from enrichment cultures on rich and poor solid agar media

866

A. Abeliovich

CHAPTER 3.2.17

Table 1. Four culture mediaa for lithoautotrophic and mixotrophic growth of nitrite oxidizers. Ingredient Distilled water (ml) Seawater (ml) NaNO2 (mg) KNO2 (mg) MgSO4 ·7H2O (mg) CaCl2 ·2H2O (mg) CaCO3 (mg) KH2PO4 (mg) K2HPO4 (mg) FeSO4 (mg) Chelated ironb (mg) Na2Mo4 (µg) (NH4)6Mo7O24 ·4H2O (µg) MnCl2 ·4H2O (µg) CoCl2 ·6H2O (µg) CuSO4 ·5H2O (µg) ZnSO4 ·7H2O (µg) KHCO3 (mg) NaCl (mg) Sodium pyruvate (mg) Yeast extractc (mg) Peptonec (mg)

1a

2b

3b

1,000

1,000

1,000

2,000

2,000

50

50

3.0 150

3.0 150

300 187.5 12.5 500 500 10

4c 300 700 69 100 6.0

1.74 0.15

0.15 1.0 30

50

50 66 0.6 6.0 30

1,500 500

500 550 1,500 1,500

a

pH is adjusted to 7.4–7.8. 13%, Geigy. c Difco. b

can check contamination by heterotrophs. It may take a year to obtain pure cultures with the technique described above. Medium 1a for terrestrial strains is from Aleem and Alexander (1958), media 2b and 3b also for terrestrial strains are from Bock and Stackebrandt (1983), and medium 4c for marine strains is from Watson and Waterbury (1971).

Habitats As described earlier, the nitrite-oxidizing bacteria are ubiquitous, and are usually found in association with the ammonia-oxidizing bacteria, which provide them with their energy source. In spite of many reports describing their sensitivity to light (Guerrero and Jones, 1996; Guerrero and Jones, 1997) and various toxicants (Deni and Penninckx, 1999; Jenicek et al., 1996), nitriteoxidizing bacteria are abundant in practically all terrestrial, marine and fresh water habitats. Nitrifying bacteria play a significant role in erosion of rocks (Lebedeva et al., 1978) primarily because they release nitrates, which dissolve carbonates, in exchange for CO2 they assimilate. This process was found to be a major cause of deterioration of historic monuments in Europe (Mansch and Bock, 1998). Proliferation of nitri-

fiers in sewage is also the basis for nitrogen removal from domestic and industrial waste in water treatment plants, including chemical industry wastewater. Nitrite oxidizers grow within a wide range of pH values up to 10 (Sorokin, 1998a; Sorokin et al., 1998b) and salinities (limits are difficult to determine as nitrifying bacteria resist very low water activities, such as occur when living within desiccated rocks). Although nitrite oxidizers are considered obligate aerobes (even though many can utilize nitrate in absence of oxygen as an electron acceptor), growth of nitrite oxidizers can be observed in totally anaerobic environments such as deep in the mud at the bottom of deep anaerobic wastewater reservoirs, in the absence of nitrate or oxygen, in presence of significant concentrations of sulfides (Abeliovich, 1987). Also, nitrifying bacteria were isolated from a marine anaerobic sediment at depths of up to 8 cm (Blackburn, 1983). Nothing is at present known about the metabolism and survival mechanisms of these nitrite-oxidizing bacteria living under strict anaerobic, reducing conditions. Light (particularly blue light) is frequently mentioned as an inhibiting factor for the nitrification process. However, the reports on the effects of light are sometimes contradictory. Nitrifiers are found in all habitats exposed to sunlight, either at the surface of fresh water or marine habitats, so obviously they must have efficient protection mechanisms to overcome photoinhibition and damage caused by direct sunlight. Sorokin et al. (1998b) and Sorokin (1998a) have isolated five strains of lithotrophic, nitriteoxidizing bacteria from sediments of three soda lakes after enrichment at pH 10 with nitrite as sole energy source; these strains were described as a new species of the genus Nitrobacter, N. alkalicus. Nitrite oxidation to nitrate occurred at pH 10. The nitrifiers had pear-shaped budding cells morphologically similar to those of Nitrobacter and formed tiny colonies on mineral nitrite agar at pH 10. In a study aimed at evaluating the potential of wastewater treatment plants to contaminate receiving waterways, survival of Nitrobacter cells associated with particles in water treatment plant discharges was studied using immunofluoresence methods (Bonnet et al., 1997). It was found that Nitrobacter colonies can settle in freshwater sediments in a week (106 cells per gram of dry sediment) and can therefore colonize river sediments. Chemolithotrophic nitrifying bacteria are dependent on the presence of oxygen for the oxidation of ammonium via nitrite to nitrate. The success of nitrification in oxygen-limited environments largely depends on the oxygen sequestering abilities of both ammonium- and

CHAPTER 3.2.17

nitrite-oxidizing bacteria. Oxygen consumption kinetics were determined with cells grown in mixed culture in chemostats at different growth rates and oxygen tensions (Laanbroek and Gerards, 1993). Reduction of oxygen tension in the culture was found to repress the oxidation of nitrite before the oxidation of ammonium was affected, and hence nitrite accumulated. In addition to the competition between ammonium- and nitrite-oxidizing bacteria, there is competition with organotrophic bacteria for the available oxygen as well. The outcome of the competition is determined by their specific affinities for oxygen as well as by their population sizes. The effect of mixotrophic growth of nitriteoxidizing Nitrobacter hamburgensis on the competition for limiting amounts of oxygen was studied in mixed continuous culture experiments with the ammonium-oxidizing Nitrosomonas europaea at different oxygen concentrations (Laanbroek et al., 1994). The specific affinity for oxygen of N. europaea was in general higher than that of N. hamburgensis, and in transient state experiments, when oxic conditions were switched to anoxic, N. hamburgensis was washed out and nitrite accumulated. However, at low oxygen concentration, the specific affinity for oxygen of N. hamburgensis increased and became as great as that of N. europaea, and owing to its larger population size, the nitriteoxidizing bacterium became the better competitor for oxygen and ammonium accumulated. Therefore Laanbroek et al. suggest that continuously oxygen-limited environments present a suitable ecological niche for the nitrite-oxidizing N. hamburgensis.

Biofilms Biofilms present a unique biotope for using molecular DNA techniques as well as FISH and various sensors. The combination of microsensor and molecular techniques is highly useful for studies on the microbial ecology of biofilms in general, and in particular for the identification of activity sites of nitrifying bacteria. Thus, the distribution of nitrifying bacteria of the genera Nitrosomonas, Nitrosospira, Nitrobacter and Nitrospina was studied by Schramm et al. (2000) in a membrane-bound biofilm system in which gradients of oxygen, pH, nitrite and nitrate were determined by means of microsensors, while the nitrifying populations along these gradients were identified and quantified using FISH in combination with confocal laser scanning microscopy. It was found that the oxic part of the biofilm was dominated by ammonia oxidizers and by members of the genus Nitrobacter, and Nitrospira sp. were virtually absent in this

The Nitrite-Oxidizing Bacteria

867

part of the biofilm, whereas they were most abundant at the oxic-anoxic interface. In the totally anoxic part of the biofilm, cell numbers of all nitrifiers were relatively low. These observations suggest the microaerophilic behavior of an as yet uncultured Nitrospira sp. as a factor affecting its environmental competitiveness. De Beer and Schramm (1999), using microsensors and molecular techniques (such as in situ hybridization with 16S rDNA-targeted oligonucleotide probes), showed that there exists in biofilms grown in bioreactors a complex nitrifying community, consisting of members of the genera Nitrosomonas, Nitrosospira, Nitrobacter and Nitrospira. In nitrifying biofilms from a trickling filter of an aquaculture water recirculation system, it was found that nitrification was restricted to a narrow zone of 50 µm of the very top of the film. Ammonia oxidizers formed a dense layer of cell clusters in the upper part of the biofilm, whereas the nitrite oxidizers showed less-dense aggregates in the proximity of the ammonia oxidizers. Both ammonia and nitrite oxidizers were not restricted to the oxic zone of the biofilm but were also detected in substantially lower numbers in the anoxic layers and even occasionally at the bottom of the biofilm (Schramm et al., 1996).

Enumeration Conventional methodologies for counting nitrifiers are gradually being replaced by new technologies, although the MPN method still dominates, this because all methods based on specific probes or antibodies still leave open the possibility that some species do not crossreact with the probe used. On the other hand, the issue of optimal conditions for maximal yields using MPN counts has not been yet resolved. One example is a new amperometric enzymelinked immunoassay for specific enumeration of Nitrobacter, which uses an electrode made of glassy carbon on which an immunological reaction is carried out. The detection limit was approximately 3 × 106 Nitrobacter cells/ml (Sanden and Dalhammar, 2000). It was shown that this method could be applied for the enumeration of Nitrobacter in activated sludge and other environmental samples. The reason for the attempts to develop new methodologies is the lengthy incubation periods required for stable MPN counts: maximum most probable numbers of the ammonia-oxidizing group were attained in 20–55 days (median, 25) and MPN estimates of nitrite oxidizers required a much longer incubation (103–113 days; Matulewich et al., 1975). In addition, Both et al. (1990a) and (1990b) studied the effect of two concentrations of nitrite in the

868

A. Abeliovich

incubation medium, 0.05 and 5.0 mM, and found that numbers of nitrite-oxidizing bacteria were highly dependent on the nitrite concentration as well as on the soil sampled. Neither the influence of pH or nitrite concentration in the incubation medium could account exclusively for the MPN-enumeration result, which raises the issue of whether to use more than one incubation medium for the enumeration of nitrite-oxidizing bacteria. Enumeration of nitrite-oxidizing bacteria in soil samples by the MPN technique often showed relatively high cell number at a low nitrite concentration, when compared to the number of ammoniumoxidizing bacteria. When different Nitrobacter species as well as nongrowing cells differing in age were incubated 5 months at 20°C in presence of various nitrite concentrations and pH values, it appeared that the growth of cells taken from an early stationary phase culture of all these species was not affected by high nitrite concentrations or low pH. Growth of 8- and 18-month-old nongrowing cells of Nitrobacter hamburgensis was also insensitive to high nitrite concentration (5 mM). The growth of 8- and 18-month old resting cells of N. vulgaris was repressed only by a combination of 5 mM nitrite and a low pH. Growth of 8-month-old nongrowing cells of N. winogradsky was sensitive to 5 mM irrespective of pH, but growth of 18-month-old cells were inhibited by 5 mM nitrite only at a low pH. The growth of 8- and 18-month-old resting cells of N. winogradsky serotype agilis was repressed by low pH rather than by high nitrite concentration (Laanbroek and Schotman, 1991). These results emphasize the problems associated with using MPN counts for estimating size of nitrifier populations in natural samples. A molecular approach based on PCR that was expected to detect and quantify nitrifying bacteria was also tested using specific primers of the genus Nitrobacter. In this study, coupled in parallel was a 14C-radiotracing method used to measure potential nitrification; it was shown that Nitrobacter represented less than 0.1% of the total bacterial community (Berthe et al., 1999). In another study, also aimed at counting Nitrobacter populations in situ by PCR, two primers from the 16S rDNA gene were used to generate a 397-bp fragment by amplification of Nitrobacter species DNA, and it was found that the PCR had a lower detection threshold (102 Nitrobacter cells per gram of soil) than did the MPN or fluorescent antibody method (Degrange and Bardin, 1995). In contrast, another study (Feray et al., 1999) found the best recovery yield was obtained with the immunofluorescence technique (21.3%), and the poorest detection level was reached with the MPN method (3.1%).

CHAPTER 3.2.17

Wastewater Tertiary wastewater treatment, aimed at removing all nitrogen wastes from the effluents, is costly in terms of the oxygen required for oxidizing ammonia to nitrate. However, as far as the nitrification—denitrification process is concerned, oxidation of nitrite to nitrate is an unnecessary step, as denitrification can just as well proceed from nitrite. Therefore, attempts are being made to devise operational parameters that will inhibit the activity of nitrite-oxidizing bacteria in wastewater treatment plants. Thus, it was found (Rhee et al., 1997) that accumulation of nitrite occurred during the aerobic phase of a sequencing batch reactor (SBR) operating to remove nitrogen from synthetic wastewater: the activity of autotrophic nitrite oxidizers was reduced in the SBR and the free ammonia was the main inhibitor of nitrite oxidation. It has been shown experimentally (Garrido et al., 1997) that it is possible to convert all ammonium to approximately 50% nitrate and 50% nitrite in the effluent of a biofilm air-lift suspension reactor, this with oxygen concentrations between 1 and 2 mg/liter. The ammonia- and nitrite-oxidizing bacterial populations occurring in the nitrifying activated sludge of an industrial plant treating sewage with high ammonia concentrations were also studied using in situ hybridization with a set of hierarchical 16S rDNA-targeted probes (Juretschko et al., 1998). Although a Nitrobacter strain was isolated, members of the genus Nitrobacter were not detectable in the activated sludge by in situ hybridization. A specific 16S rDNA-targeted probe for Nitrospira demonstrated that a Nitrospira-like bacterium was present in significant numbers (9% of the total bacterial counts). According to Blackall (2000), even though it is common knowledge that Nitrosomonas and Nitrobacter are the major ammonia and nitrite oxidizers, today we know that these organisms may not play any role in the transformations for which they have achieved such acclaim. Using the above-mentioned methodologies, Nitrospiralike bacteria were found to be the dominant nitrite oxidizers in both enriched and full scale nitrifying systems.

Resistance, Inhibition and Biodegradation Because inhibition of nitrite oxidation is economically advantageous in wastewater treatment plants, procedures were developed for identification of nitrification inhibitors in wastewater.

CHAPTER 3.2.17

These include fractionation of wastewater samples and a nitrification inhibition assay with pure cultures of Nitrobacter to identify the inhibitory effect. In such a study, a series of unsaturated fatty acids and two monoterpenes were found to be inhibitory in an industrial wastewater sample (Svenson et al., 2000). A shorter nitrification-denitrification cycle occurs in the presence of free ammonium. Although the occurrence of this cycle depends on both a high ammonia concentration and high pH, it was found that the pH of the wastewater is the decisive parameter (Surmacz-Gorska et al., 1997). However, a study aimed at identifying nitrification at high pH values in soda lakes and soda soils with pH 9.5–11 led to the isolation of a nitrifier morphologically similar to Nitrobacter that formed tiny colonies on mineral nitrite agar at pH 10 (Sorokin, 1998a). Hwang et al. (2000), in a detailed study of the inverse relation between alkalinity and ammonia with respect to inhibition of nitrite oxidation, showed that when the molar ratio of carbonate alkalinity to ammonia increased from 4.1 to 9.4 (thus increasing the concentration of free ammonia), the ammonium removal rate doubled. At the same time the higher concentration of free ammonia in the medium was a selective inhibitor for Nitrobacter, causing an enhanced nitrite build-up in a biofilm reactor. As for the effect of heavy metals, laboratory evaluations were conducted to study the toxic effects of copper and nickel on a culture of strictly obligatory nitrifiers (Nitrosomonas sp. and Nitrobacter sp.) in continuous flow stirred tank reactors. Nitrosomonas sp. was found to be equally or more sensitive than Nitrobacter sp. (Lee et al., 1997). Attempts were made to develop a process for the simultaneous removal of organic halogens and nitrogen from kraft pulp mill effluents. A nitrifying biofilm reactor removed organic halogens from bleached kraft pulp mill effluents including chlorophenols from synthetic wastewater (Kostyal et al., 1997; Kostyal et al., 1998). In another study, none of several chemicals tested (nonylphenol, naphthalene, 2-methylnaphthalene, di-2-ethylhexylphthalate and toluene) inhibited nitrification when added to soil at various concentrations (Kirchmann et al., 1991). However, in many cases municipal wastewater did contain substances that inhibited nitrification to varying degrees (Jonsson et al., 2000). Reports conflict on the effect of toluene on indigenous microbial populations. Nitrite oxidation potential (NOP) was reduced after incubation with high toluene concentrations for 45 days (Fuller and Scow, 1996). Trichloroethylene is also inhibitory to the soil indigenous nitrifying population (Fuller and Scow, 1997).

The Nitrite-Oxidizing Bacteria

869

Volatile fatty acids such as formic, acetic, propionic, n-butyric, isobutyric, n-valeric, isovaleric and n-caproic acid as well as trimethylamine are inhibitory to nitrification, but only at high concentrations, suggesting that volatile fatty acids and trimethylamine alone cannot account for the inhibition of the nitrification activity in domestic wastewater (Eilersen et al., 1994). Nitrifying bacteria were also found to be resistant to monochloramines, and the combination of increased concentrations of oxidized nitrogen with decreased total chlorine in treated water was used as an indicator of bacterial nitrification (Cunliffe, 1991). The need to set limits on nitrite oxidation in wastewater treatment plants created a demand for assays to determine the degree of inhibition of nitrification. One such assay, developed by Grunditz and Dalhammar, was based on inhibition of pure cultures of Nitrosomonas and Nitrobacter isolated from sewage sludge (Grunditz and Dalhammar, 2001). As for the effects of light, nitrifying bacteria (both ammonia and nitrite oxidizers) are capable of recovery from photoinhibition in the dark (see above). Recovery of oxidizing activities is both dose and wavelength dependent (Guerrero and Jones, 1996). The light-absorbing pigment was identified as a porphyrin-like pigment with an absorption maximum at 408 nm, accumulating at the late exponential phase of growth. This photoreceptor was found at higher concentrations in ammonia oxidizers than in nitrite oxidizers (Guerrero and Jones, 1997).

Lithotrophy An evaluation of field data from historical buildings in Germany, carried on by Mansch and Bock (1998), showed that ammonia and nitrite oxidizers were found in 55 and 62% of the samples, respectively, and that nitrifying bacteria will colonize natural stone within several years of exposure. The highest cell numbers were in some cases found underneath the surface. Nitrifying bacteria showed a preference for calcareous material with a pore radius of 1–10 µm. Cell numbers of nitrifying bacteria did not correlate with the nitrate content of the stone material. Their data strongly suggest that microbial colonization of historical buildings is enhanced by anthropogenic air pollution. A comparison of samples taken between 1990 and 1995 from buildings throughout Germany showed that in eastern Germany significantly more colonization with facultatively methylotrophic bacteria and nitrifying bacteria existed. The same was true for natural stone from an urban exposure site when compared to material from a rural exposure site.

870

A. Abeliovich

CHAPTER 3.2.17

Literature Cited

Eilersen, A. M., M. Henze, and L. Kloft. 1994. Effect of volatile fatty-acids and trimethylamine On nitrification in activated-sludge. Water Res. 28:1329–1336. Feray, C., B. Volat, V. Degrange, A. Clays-Josserand, and B. Montuelle. 1999. Assessment of three methods for detection and quantification of nitrite-oxidizing bacteria and nitrobacter in freshwater sediments. Microb. Ecol. 37:208–217. Fuller, M. E., and K. M. Scow. 1996. Effects of toluene on microbially-mediated processes involved in the soil nitrogen cycle. Microb. Ecol. 32:171–184. Fuller, M. E., and K. M. Scow. 1997. Impact of trichloroethylene and toluene on nitrogen cycling in soil. Appl. Environ. Microbiol. 63:4015–4019. Garrido, J. M., W. A. J. van Benthum, M. C. M. van Loosdrecht, and J. J. Heijnen. 1997. Influence of dissolved oxygen concentration on nitrite accumulation in a biofilm airlift suspension reactor. Biotechnol. Bioeng. 53:168–178. Gieseke, A., U. Purkhold, M. Wagner, R. Amann, and A. Schramm. 2001. Community structure and activity dynamics of nitrifying bacteria in a phosphate-removing biofilm. Appl. Environ. Microbiol. 67:1351–1362. Grunditz, C., and G. Dalhammar. 2001. Development of nitrification inhibition assays using pure cultures of Nitrosomonas and Nitrobacter. Water Res. 35:433–440. Guerrero, M. A., R. D. Jones. 1996. Photoinhibition of marine nitrifying bacteria. 2: Dark recovery after monochromatic or polychromatic irradiation. Marine Ecology (Progress Series) 141:193–198. Guerrero, M. A., and R. D. Jones. 1997. Light-induced absorbance changes associated with photoinhibition and pigments in nitrifying bacteria. Aquat. Microb. Ecol. 13:233–239. Guschin, D. Y., B. K. Mobarry, D. Proudnikov, D. A. Stahl, B. E. Rittmann, and A. D. Mirzabekov. 1997. Oligonucleotide microchips as genosensors for determinative and environmental studies in microbiology. Appl. Environ. Microbiol. 63:2397–2402. Hovanec, T. A., L. T. Taylor, A. Blakis, and E. F. Delong. 1998. Nitrospira-like bacteria associated with nitrite oxidation in freshwater aquaria. Appl. Environ. Microbiol. 64: 258–264. Hwang, B. H., K. Y. Hwang, E. S. Choi, D. K. Choi, and J. Y. Jung. 2000. Enhanced nitrite build-up in proportion to increasing alklinity/NH4+ ratio of influent in biofilm reactor. Biotchnol. Lett. 22:1287–1290. Jenicek, P., J. Zabranska, and M. Dohanyos. 1996. The influence of anaerobic pretreatment on the nitrogen removal from biosynthetic pharmaceutical wastewaters. Ant. v. Leeuwenhoek 69:41–46. Jonsson, K., C. Grunditz, G. Dalhammar, and J. L. Jansen. 2000. Occurrence of nitrification inhibition in Swedish municipal wastewaters. Water Res. 34:2455–2462. Juretschko, S., G. Timmermann, M. Schmid, K. H. Schleifer, A. Pommerening-Roser, H. P. Koops, and M. Wagner. 1998. Combined molecular and conventional analyses of nitrifying bacterium diversity in activated sludge: Nitrosococcus mobilis and Nitrospira-like bacteria as dominant populations. Appl. Environ. Microbiol. 64:3042–3051. Kirchmann, H., H. Astrom, and G. Jonsall. 1991. Organic pollutants in sewage-sludge. 1: Effect of toluene, naphthale, 2-methylnaphthalene, 4-normal-nonylphenol and di-2-ethylhexyl phthalate on soil biological processes and their decomposition in soil. Swedish J. Agric. Res. 21:107–113.

Abeliovich, A. 1987. Nitrifying bacteria in wastewater reservoirs. Appl. Environ. Microbiol. 53:754–760. Aleem, M. I. H., and M. Alexander. 1958. Cell free nitrification by nitrobacter. J. Bacteriol. 76:510–514. Bartosch, S., I. Wolgast, E. Spieck, and E. Bock. 1999. Identification of nitrite-oxidizing bacteria with monoclonal antibodies recognizing the nitrite oxidoreductase. Appl. Environ. Microbiol. 65:4126–4133. Berthe, T., J. Garnier, and F. Petit. 1999. Quantification of nitrifying bacteria of the genus Nitrobacter in an aquatic system (Seine estuary, France). Science 322:517–526. Blackall, L. L. 2000. A summary of recent microbial discoveries in biological nutrient removal from wastewater. Austral. Biotechnol. 10:29–31. Blackburn, H. 1983. The microbial nitrogen cycle. In: W. E. Krumbein (Ed.) Microbial Geochemistry. Blackwell Scientific Publications. Boston, MA. 63–89. Bock, E., H. Sundermeyer-Klinger, and E. Stackebrandt. 1983. New facultative lithoautotrophic nitrite oxidizing bacteria. Arch. Microbiol. 136:281–284. Bock, E., and H. P. Koops. 1992. The genus nitrobacter and related genera. In: A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K. H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer-Verlag. New York, NY. 3:2302–2309. Bonnet, C., B. Volat, R. Bardin, V. Degranges, and B. Montuelle. 1997. Use of immunofluorescence technique for studying a Nitrobacter population from wastewater treatment plant following discharge in river sediments: First experimental data. Water Res. 31:661–664. Both, G. J., S. Gerards, and H. J. Laanbroek. 1990a. Enumeration of nitrite oxidizing bacteria in grassland soils using a most probable technique: Effect of nitrite concentration and sampling procedure. FEMS Microbiol. Ecol. 74:277–285. Both, G. J., S. Gerards, and H. J. Laanbroek. 1990b. Most probable numbers of chemlitho-autotrophic nitriteoxidizing bacteria in well drained grassland soils: Stimulation by high nitrite concentrations. FEMS Microbiol. Ecol. 74:287–293. Burrell, P. C., J. Keller, and L. L. Blackall. 1998. Microbiology of a nitrite-oxidizing bioreactor. Appl. Environ. Microbiol. 64:1878–1883. Burrell, P., J. Keller, and L. L. Blackall. 1999. Characterisation of the bacterial consortium involved in nitrite oxidation in activated sludge. Water Sci. Technol. 39:45–52. Cunliffe, D. A. 1991. Bacterial nitrification in chloraminated water supplies. Appl. Environ. Microbiol. 57:3399– 3402. de Beer, D., and A. Schramm. 1999. Micro-environments and mass transfer phenomena in biofilms studied with microsensors. Water Sci. Technol. 39:173–178. Degrange, V., and R. Bardin. 1995. Detection and counting of Nitrobacter populations in soil by PCR. Appl. Environ. Microbiol. 61:2093–2098. Deni, J., and M. J. Penninckx. 1999. Nitrification and autotrophic nitrifying bacteria in a hydrocarbonpolluted soil. Appl. Environ. Microbiol. 65:4008–4013. Ehrich, S., D. Behrens, E. Lebedeva, W. Ludwig, and E. Bock. 1995. A new obligately chemolithoautotrophic, nitriteoxidizing bacterium, Nitrospira moscoviensis sp. nov. and its phylogenetic relationship. Arch. Microbiol. 164:16–23.

CHAPTER 3.2.17 Kirstein, K., and E. Bock. 1993. Close genetic relationship between Nitrobacter hamburgensis nitrite oxidoreductase and Escherichia coli nitrate reductases. Arch. Microbiol. 160:447–453. Kostyal, E., E. L. Nurmiaho Lassila, J. A. Puhakka, and M. Salkinoja Salonen. 1997. Nitrification, denitrification, and dechlorination in bleached kraft pulp mill wastewater. Appl. Microbiol. Biotechnol. 47:734–741. Kostyal, E., M. Borsanyi, L. Rigottier-Gois, and M. S. Salkinoja-Salonen. 1998. Organic halogen removal from chlorinated humic ground water and lake water by nitrifying fluidized-bed biomass characterised by electron microscopy and molecular methods. Appl. Microbiol. Biotechnol. 50:612–622. Laanbroek, H. J., and J. M. T. Schotman. 1991. Effect of nitrite concentration and pH on most probable number enumerations of non growing nitrobacter spp. FEMS Microbiol. Ecol. 85:269–278. Laanbroek, H. J., and S. Gerards. 1993. Competition for limiting amounts of oxygen between Nitrosomonas europaea and Nitrobacter winogradskyi grown in mixed continuous cultures. Arch. Microbiol. 159:453–459. Laanbroek, H. J., P. L. E. Bodelier, and S. Gerards. 1994. Oxygen consumption kinetics of Nitrosomonas europaea and Nitrobacter hamburgensis in mixed continuous cultures at different oxygen concentrations. Arch. Microbiol. 161:156–162. Lebedeva E. V., N. N. Lialikova, and I. Bugel’skii. 1978. Participation of nitrifying bacteria in the disintegration of serpentinous ultrabasic rock. Mikrobiologiia 47:1101– 1107. Lee, Y. W., S. K. Ong, and C. Sato. 1997. Effects of heavy metals on nitrifying bacteria. Water Sci. Technol. 36:69– 74. Macdonald, R. M. 1986. Nitrification in soil: An introductory history. In: J. I. Prosser (Ed.) Nitrification. IRL Press. Washington, DC, Washington, DC. 20:1–17. Mansch, R., and E. Bock. 1998. Biodeterioration of natural stone with special reference to nitrifying bacteria. Biodegradation 9:47–64. Matulewich, V. A., P. F. Strom, and M. S. Finstein. 1975. Length of incubation for enumerating nitrifying bacteria present in various environments. Appl. Microbiol. 29:265–268. Meincke, M., E. Bock, D. Kastrau, and P. M. H. Kroneck. 1992. Nitrite oxidoreductase from Nitrobacterhamburgensis-redox centers and their catalytic role. Arch. Microbiol. 158:127–131. Mobarry, B. K., M. Wagner, V. Urbain, B. E. Rittmann, and D. A. Stahl. 1996. Phylogenetic probes for analyzing abundance and spatial organization of nitrifying bacteria. Appl. Environ. Microbiol. 62:2156–2162. Navarro, E., M. P. Fernandez, F. Grimont, A. Claysjosserand, and R. Bardin. 1992a. Genomic heterogeneity of the genus Nitrobacter. Int. J. Syst. Bacteriol. 42:554–560. Navarro, E., P. Simonet, P. Normand, and R. Bardin. 1992b. Characterization of natural populations of Nitrobacter spp. using PCR/RFLP analysis of the ribosomal intergenic spacer. Arch. Microbiol. 157:107–115. Noda, N., H. Ikuta, Y. Ebie, A. Hirata, S. Tsuneda, M. Matsumura, T. Sumino, and Y. Inamori. 2000. Rapid quantification and in situ detection of nitrifying bacteria in biofilms by monoclonal antibody method. Water Sci. Technol. 41:301–308. Okabe, S., H. Satoh, and Y. Watanabe. 1999. In situ analysis of nitrifying biofilms as determined by in situ hybridiza-

The Nitrite-Oxidizing Bacteria

871

tion and the use of microelectrodes. Appl. Environ. Microbiol. 65:3182–3191. Orso, S., M. Gouy, E. Navarro, and P. Normand. 1994. Molecular phylogenetic analysis of Nitrobacter spp. Int. J. Syst. Bacteriol. 44:83–86. Pasteur, L. 1862. Etudes sur les mycoderms: Rôle de ces plantes dans la fermentation acétique. C. R. Acad. Sci. 54:265–270. Rhee, S. K., J. J. Lee, and S. T. Lee. 1997. Nitrite accumulation in a sequencing batch reactor during the aerobic phase of biological nitrogen removal. Biotechnol. Lett. 19:195– 198. Sakai, K., Y. Ikehata, Y. Ikenaga, M. Wakayama, and M. Moriguchi. 1996. Nitrite oxidation by heterotrophic bacteria under various nutritional and aerobic conditions. J. Ferment. Bioeng. 82:613–617. Sakai, K., K. Nakamura, M. Wakayama, and M. Moriguchi. 1997. Change in nitrite conversion direction from oxidation to reduction in heterotrophic bacteria depending on the aeration conditions. J. Ferment. Bioeng. 84:47–52. Sanden B., and G. Dalhammar. 2000. Application of an amperometric immuno sensor for the enumeration of Nitrobacter in activated sludge. Appl. Microbiol. Biotechnol. 54:413–417. Schramm, A., L. H. Larsen, N. P. Revsbech, N. B. Ramsing, R. Amann, and K. H. Schleifer. 1996. Structure and function of a nitrifying biofilm as determined by in situ hybridization and the use of microelectrodes. Appl. Environ. Microbiol. 62:4641–4647. Schramm, A., D. De Beer, M. Wagner, and R. Amann. 1998. Identification and activities in situ of Nitrosospira and Nitrospira spp. as dominant populations in a nitrifying fluidized bed reactor. Appl. Environ. Microbiol. 64: 3480–3485. Schramm, A., D. De Beer, A. Gieseke, and R. Amann. 2000. Microenvironments and distribution of nitrifying bacteria in a membrane-bound biofilm. Environ. Microbiol. 2:680–686. Sorokin, D. Y. 1991. Peroxide mechanism of nitrite oxidation in heterotrophic bacteria. Microbiology 60:4–9. Sorokin, D. Y. 1998a. On the possibility of nitrification in extremely alkaline soda biotopes. Microbiology 67:335– 339. Sorokin, D. Y., G. Muyzer, T. Brinkhoff, J. G. Kuenen, and M. S. Jetten. 1998b. Isolation and characterization of a novel facultatively alkaliphilic Nitrobacter species, N. alkalicus sp. nov. Arch. Microbiol. 170: 345–352. Surmacz-Gorska, J., A. Cichon, and K. Miksch. 1997. Nitrogen removal from wastewater with high ammonia nitrogen concentration via shorter nitrification and denitrification. Water Sci. Technol. 36:73–78. Svenson, A., B. Sanden, G. Dalhammar, M. Remberger, and L. Kaj. 2000. Toxicity identification and evaluation of nitrification inhibitors in wastewaters. Environ. Toxicol. 15:527–532. Teske, A., E. Alm, J. M. Regan, S. Toze, B. E. Rittmann, and D. A. Stahl. 1994. Evolutionary relationships among ammonia- and nitrite-oxidizing bacteria. J. Bacteriol. 176:6623–6630. Watson, S. W., and J. B. Waterbury. 1971. Characteristics of two marine nitrite oxidizing bacteria, Nitrospina gracilis nov. gen. nov. sp. and Nitrococcus mobilis nov. gen. nov. sp. Arch. Mikrobiol. 77:203–230. Watson, S. W., F. W. Valois, and J. B. Waterbury. 1981. The family Nitrobacteraceae. In: M. P. Starr, H. Stolp, H. G. Trüper, A. Balows, and H. G. Schlegel (Eds.) {The

872

A. Abeliovich

Prokaryotes}. Springer-Verlag. New York, NY. 1:1005– 1022. Watson, S. W., E. Bock, F. W. Valois, J. B. Waterbury, and U. Schlosser. 1986. Nitrospira marina gen. nov., sp. nov.: A chemolithotophic nitrite oxidizing bacterium. Arch. Microbiol. 144:1–7. Woese, C. R., W. G. Weisburg, B. J. Paster, C. M. Hahn, R. S. Tanner, N. R. Krieg, H. P. Koops, H. Harms, and

CHAPTER 3.2.17 E. Stackebrandt. 1984. The phylogeny of purple bacteria: The beta subdivision. Syst. Appl. Microbiol. 5:327–336. Woese, C. R., W. G. Weisburg, C. M. Hahn, B. J. Paster, L. B. Zablen, B. J. Lewis, T. J. Macke, W. Ludwig, and E. Stackebrandt. 1985. The phylogeny of purple bacteria: The gamma subdivision. Syst. Appl. Microbiol. 6:25– 33.

Prokaryotes (2006) 5:873–891 DOI: 10.1007/0-387-30745-1_42

CHAPTER 3.2.18 ehT

areneG

, suc raozA

, o i rb i vozA

ar i psozA

dna

The Genera Azoarcus, Azovibrio, Azospira and Azonexus BARBARA REINHOLD-HUREK AND THOMAS HUREK

Introduction The genera Azoarcus, Azospira, Azovibrio and Azonexus harbor mostly nitrogen-fixing Betaproteobacteria. In their first taxonomic description, several strains which were later assigned to the genera Azovibrio, Azospira (Reinhold-Hurek et al., 1993) and Azonexus (Hurek et al., 1997b) were included in the genus Azoarcus sensu lato. Therefore, they are discussed in one chapter. Except for one strain, all these diazotrophic bacteria had been isolated from a similar source, roots of a pioneer grass in Pakistan or fungal resting stages from the same field; they had many physiological features in common, and they were phylogenetically related. Therefore, three groups of bacteria were included into Azoarcus although they were located on the Azoarcus rRNA branch at low Tm(e) values (Reinhold-Hurek et al., 1993) or showed a low 16S rDNA similarity (Hurek et al., 1997b). Isolation of additional strains, which made possible a better phenotypic description, and sequencing of almost complete 16S rDNA sequences allowed the reassessment of the taxonomic structure of Azoarcus sensu lato. The unnamed groups C and D (Reinhold-Hurek et al., 1993) obtained the rank of different genera Azovibrio and Azospira, respectively (ReinholdHurek and Hurek, 2000); Azoarcus sensu lato group E (Hurek et al., 1997b) was proposed as Azonexus (Hurek et al., 1997b). Strains of these genera deserve special recognition as endophytes of grass roots or as degraders of aromatic compounds.

The Genus Azoarcus Introduction The genus Azoarcus harbors two ecologically different groups of bacteria: 1) soil-borne strains which can degrade aromatic hydrocarbons under denitrifying conditions, and 2) bacteria associated with grass roots epi- or endophytically, which apparently do not survive well in root-free soil. They will be referred to as “soil-borne” or

“plant-associated” species, respectively, in this chapter. Azoarcus was originally described with two valid species of grass-associated diazotrophs (Reinhold-Hurek et al., 1993). However, a growing number of species of soil-borne strains has been added in recent years.

Phylogeny Azoarcus spp. are members of the Betaproteobacteria in the Rhodocyclales according to phylogenetic analysis of almost complete 16S rRNA gene sequences. They are phylogenetically most closely related to Thauera with which they form the Azoarcus/Thauera branch (Fig. 1). Azoarcus species are located on two different clades, which in part reflects their physiology and ecology: the soil-borne species A. tolulyticus, A. toluclasticus, A. toluvorans, A. evansii, A. buckelii and A. anaerobius and the plant-associated species A. indigens, A. communis and Azoarcus sp. strain BH72. The analysis of phylogenetic relationships within the Azoarcus/Thauera 16S rDNA cluster is rendered difficult because the branching pattern between Thauera, the soil-borne and the plant-associated Azoarcus species is unstable when tested with different tree-building methods. This was also observed previously (Hurek et al., 1997b; Reinhold-Hurek and Hurek, 2000). Also in other taxa closely related to each other, like Rhizobium and related genera, the resolution of phylogenetic analysis is sometimes limited. Nevertheless, all named species in the genus Thauera cluster in one clade with highly significant support levels (Fig. 1), allowing a reliable assignment to this genus. Also most soil-borne species or strains of Azoarcus fall into one clade with significant support levels, and none of the species containing plant-associated strains are located on this branch. The 16S rDNA phylogenetic distances within these three clades are similar. Within the genus Azoarcus, they are up to 6%, while Thauera is 6–7% distant. Thus Azoarcus spp. represents a rather heterogeneous group. Moreover, since the phylogenetic distances within the three clades in the Azoarcus/ Thauera group are similar, both subgroups of

suxenozA

874

B. Rareinhold-Hurek and T. Hurek

CHAPTER 3.2.18 Azoarcus sp. Td-19 (L33690) Azoarcus sp. Td-3 (L33693) Azoarcus sp. ToN1 (X83534) 75 Azoarcus evansii pF6. (U44853) Azoarcus evansii KB740 r (X77679) 92 26 Azoarcus toluvorans Td-21 r (L33692) Azoarcus sp. M3 (Y11040) 52 37 Azoarcus sp. Td-15 (L33688) 83 Azoarcus sp. T (AF129465) 89 Azoarcus toluvorans Td-17 (L33689) 44 Azoarcus sp. CC-11 (AB033745) 68 Azoarcus sp. FL05 (AF011330) Azoarcus toluclasticus MF7 (AF123076) 50 Azoarcus toluclasticus MF63 r (AF123077) 96 Azoarcus tolulyticus Td-2 (L33691) 61 84 Azoarcus tolulyticus BL11 (AF123075) 56 Azoarcus tolulyticus Td-1 (L33687) 53 Azoarcus tolulyticus Tol-4 r (L33694) Azoarcus sp. 22Lin (Y13222) Azoarcus sp. T3 (Y11041) Azoarcus sp. CR23 (AF011328) 62 Azoarcus sp. EbN1 (X83531) 99 Azoarcus buckelii U120 (AJ315676) Azoarcus anaerobius LuFRes1 r (Y14701) 74 Azoarcus sp. HxN1 (AF331975) 38 Azoarcus sp. PBN1 (X83532) 88 71

43 44

47 97

55

71

25

52 100 63 100 52 80

96 100

70

50

43 63 44

38

32

48 20 23

Uncultured bacterium clone A33 (AF072925) Uncultured bacterium clone H30 (AF072924) Uncultured bacterium clone H25 (AF072919) 100 Uncultured bacterium clone S23 (AF072921) 85 83 Uncultured bacterium clone S3 (AF072918) 100 Azoarcus indigens ME14 Azoarcus indigens VB32 r (AF011345) Azoarcus sp. BH72 (AF011344) Azoarcus sp. LU1 (AJ007007) Azoarcus communis SWub3 r (AF011343) 95 Azoarcus communis ME91 Uncultured bacterium clone OS ac16 Thauera

Duganella 98 Azovibrio restrictus S5b2 r (AF011346) Azovibrio sp. BS20-3 (AF011349) Uncultured bacterium clone SJA-10 (AJ009452) Azonexus fungiphilus BS5-8 r (AF011350) Dechloromonas sp. MissR (AF170357) 66 48 Dechloromonas sp. SIUL (AF170356) 55 Dechloromonas sp. JJ (AY032611) 72 Ferribacterium limneticum cda-1 (Y17060) 46 Isolate OcN1 (AF331976) Isolate B2 (AF035048) Uncultured bacterium clone SBRH147 (AF204246) 100 25 Dechloromonas agitatus CBK (AF047462) 44 100 Dechloromonas sp. CL (AF170354) Dechloromonas sp. NM (AF170355) 86 Uncultured bacterium clone SA35 (AF245350) Uncultured bacterium clone PHOS-HE23 (AF314420) Isolate B5 (AF035051) 100 Isolate 51885 (AF227840) Oral isolate A08KA (AY005031) Rhodocyclus tenuis DSM 110 (D16209) Rhodocyclus purpureus DSM 168 (M34132) 96 Uncultured bacterium clone SJA-109 (AJ009484) 97 Propionivibrio dicarboxylicus DSM 5885 (Y17601) Propionibacter pelophilus asp 66 (AF016690) Azospira oryzae 6a3 (AF011347) 96 Uncultured bacterium clone SJA-52 (AJ009466) Dechlorosoma] sp. SDGM (AF170349) 100 64 Dechlorosoma] sp. Iso2 (AF170351) 51 Dechlorosoma] sp. Iso1 (AF170350) 48 Dechlorosoma suillum] ps (AF170348)

0.01

Fig. 1. Phylogenetic analysis of 16S rDNA sequences (1358 positions) of a subgroup of the Betaproteobacteria. Subtree of the Rhodocyclus/Thauera/Azoarcus group derived from an analysis of 159 sequences. Tree inference was carried out using the neighbor-joining algorithm and a Jukes-Cantor correction with 125 bootstrap repetitions. Sequence accession numbers are given in parentheses.

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Azoarcus might also deserve the rank of different genera in the future. However, this will require a rigid polyphasic taxonomic analysis. On the basis of 16S rDNA sequence analyses, all of the validly described species of Azoarcus are well resolved except for the two strains of A. toluvorans, which are interdispersed with several strains of uncertain affiliation (Fig. 1). Former members of A. tolulyticus, Td-3 and Td-19, have recently been removed from this species owing to low DNA-DNA similarity (below 40%), which is also reflected in our phylogenetic analysis (Fig. 1). Interestingly, also a deeply-branching clade of as yet uncultured bacteria detected in nitrifyingdenitrifying activated sludge of an industrial wastewater treatment plant (Juretschko et al., 2002) is localized within the Azoarcus cluster (Fig. 1).

875

and Hurek, 2000; see also the section on Identification in this Chapter). Therefore, both subgroups of Azoarcus might deserve the rank of different genera in future. However, this will require a rigid polyphasic taxonomic analysis. As can be seen from the descriptive table (Table 3), many features have not been tested for all species or strains. There appears to be a bias in testing, e.g., nitrogen fixation and aerobic nutritional profiles are not excessively tested in soil-borne strains, whereas nutritional profiles under conditions of denitrification are missing for plantassociated species. These studies should also include the numerous soil-borne isolates not yet assigned to any species. As some strains are deeply branching in the phylogenetic analysis (e.g., 22Lin, T3 and M3; Fig. 1), the description of new species might be expected.

Habitat Taxonomy The definition of the genus Azoarcus is quite complex. It includes strains with a variety of physiological and ecological properties, which place them into at least two groups. In addition to their ecology, the species containing plantassociated strains differ from the soil-borne species in some nutritional features, e.g., in their inability to use carbohydrates as sole carbon source or, under conditions of denitrification, a wide range of aromatic compounds. The genus Azoarcus consists of the following eight validly described species, mostly based on polyphasic taxonomic approaches including DNA-DNA hybridization, protein profiles, fatty acid analysis, and nutritional profiles: A. indigens (type species) and A. communis (Reinhold-Hurek et al., 1993), A. tolulyticus (Zhou et al., 1995), A. toluclasticus and A. toluvorans (Song et al., 1999), A. evansii (Anders et al., 1995), A. anaerobius (Springer et al., 1998) and A. buckelii (Mechichi et al., 2002). One isolate (BH72) is different from these at the species level according to DNADNA similarity studies (≤25% DNA binding); however, a species name was not given because it is only a singular strain (Reinhold-Hurek et al., 1993). Numerous additional soil-borne strains are localized in the Azoarcus clade according to 16S rDNA sequence analysis (examples given in Fig. 1). Most have been isolated under conditions of denitrification with aromatic hydrocarbons as a C-source, but the lack of nutritional/physiological data and DNA-DNA similarity values does not yet allow a species assignment. The phylogenetic distances within the three clades in the Azoarcus/Thauera group are similar (Reinhold-Hurek and Hurek, 2000), and the soilborne and plant-associated species show considerable phenotypic differences (Reinhold-Hurek

Azoarcus spp. occur in terrestrial ecosystems; however, both groups of Azoarcus spp. strongly differ in their habitats and ecology. Members of A. indigens, A. communis and Azoarcus sp. BH72 occur inside roots or on the root surface of Gramineae and have never been isolated from root-free soil so far (Reinhold-Hurek and Hurek, 1998b), except for members of A. communis originating not from plants but from French refinery oily sludge (S2; Laguerre et al., 1987; Reinhold-Hurek et al., 1993) or from a compost biofilter in Canada (Lu1; Juteau et al., 1999). In contrast, all isolates localized in the clade of A. tolulyticus, A. toluvorans, A. toluclasticus, A. evansii, A. buckelii or A. anaerobius do not originate from living plants but mostly from soil and sediments. Therefore, their habitats will be treated separately. The plant-associated strains were originally isolated from Kallar grass (Leptochloa fusca [L.] Kunth), a flood-tolerant salt marsh grass grown as a pioneer plant on salt-affected, flooded, lowfertility soils in the Punjab of Pakistan since the 1970s (Reinhold et al., 1986; Reinhold-Hurek et al., 1993). These nitrogen-fixing bacteria were found in high numbers in surface-sterilized roots or culms and rarely on the root surface (A. communis). Only much later, the host range was found to be extended to rice (Oryza sativa) from Nepal (Engelhard et al., 2000) or to resting stages (sclerotia) of a plant-associated basidiomycete found in rice field soil from Pakistan (Hurek et al., 1997b). Soil-borne species of Azoarcus are very widespread but not plant-associated. Strains belonging to the validly described species have been isolated from noncontaminated soils (Song et al., 1999) or contaminated soils, either with unknown contaminations in industrial areas

876

B. Rareinhold-Hurek and T. Hurek

(Song et al., 1999) or with petroleum contaminations (Fries et al., 1994; Zhou et al., 1995). Many strains were also cultured from sediments of creeks (uncontaminated or contaminated; Anders et al., 1995), aquifers (Fries et al., 1994; Fries et al., 1997; Zhou et al., 1995), or even from activated sewage sludge (Springer et al., 1998). Soil-borne Azoarcus spp. are also widespread with respect to their geographical distribution: they have been found for example in North America (United States, Canada and Puerto Rico), South America (Brazil), or Europe (Germany and Switzerland).

Isolation Plant-associated and soil-borne members of Azoarcus spp. will be treated separately. Plantassociated strains are best enriched on media free of combined nitrogen. Washed or surfacesterilized roots are macerated aseptically in wash medium (enrichment medium free of a carbon source). To avoid overgrowth by faster growing diazotrophs, inoculate enrichment cultures with serial dilutions of this material. N-free semisolid synthetic malate medium (SM-N) is used for enrichment, composed of (per liter of distilled water): MgSO4 · 2H2O, 0.2 g; NaCl, 0.1 g; CaCl2, 0.02 g; MnSO4 · H2O, 0.01 g; Na2MoO4 · 2H2O, 0.002 g; Fe(III)-EDTA, 0.066 g; KH2PO4, 0.6 g, and K2HPO4, 0.4 g, adjusted to pH 6.8; DL-malic acid, 5 g, and KOH, 4.5 g adjusted to pH 6.8; agar, 2 g; and vitamin solution, 1 ml. The vitamin solution contains (mg per liter of distilled water) myo-inositol, 10,000; niacinamide, 100; pyridoxine HCl, 100; thiamine HCl, 100; calcium pantothenate, 50; folic acid, 20; choline chloride, 50; riboflavin, 10; ascorbic acid, 100; p-aminobenzoic acid, 1; vitamin A, 0.5; vitamin D3, 0.5; vitamin B12, 0.5; and D-biotin, 0.5. Add phosphate buffer and vitamin solution after autoclaving. Inoculate medium solidified in screw-cap tubes with 10 µl of macerate or root pieces below the medium surface. Incubate tubes without shaking at 30°C or 37°C, and check for development of a subsurface pellicle. Take samples for streaking before the pellicle has moved to the medium surface and become very dense. Isolate single colonies on SM-medium agar supplemented with 20 mg of yeast extract per liter. Use acetylene reduction method to analyze enrichment cultures or isolates for nitrogenase activity in semisolid medium. Check purity and colony color on complex medium, VM ethanol (which is based on SM medium but supplemented [per liter] with “Lab Lemco” powder [Oxoid] or Bacto Peptone [Difco], 3 g; yeast extract, 1 g; NaCl, 1 g; NH4Cl, 0.5 g; and agar, 15 g). Alternatively, for saltaffected habitats, use SSM medium which is

CHAPTER 3.2.18

adapted to high salt concentrations of salinesodic soil (Reinhold et al., 1986) instead of SM medium for enrichment and isolation. For soil-borne strains, enrichment is routinely done under strictly anaerobic conditions, with nitrate as terminal electron acceptor. A variety of aromatic carbon sources have been used, depending on the species or strain under study. Preparation of media and cultivation of bacteria have to be carried out under strictly anoxic conditions. Use a typical medium (for A. evansii), which consists of (per liter of distilled water): KH2PO4, 0.816 g; K2HPO4, 5.920 g; NH4Cl, 0.530 g; MgSO4 · 7H2O, 0.200 g; KNO3, 0.5 g; and CaCl2 · 2H2O, 0.025 g; dissolve phosphate and other ingredients separately from each other, adjust both to pH 7.8 for A. evansii or neutral pH for other strains, and combine both autoclaved solutions after cooling. Add 10 ml of sterile trace elements SL-10 and 5 ml of vitamin solution. SL-1 contains (per liter of distilled water): HCl (25%; 7.7 M), 10 ml; FeCl2·4H2O, 1.5 g; ZnCl2, 70 mg; MnCl2 · 4H2O, 100 mg; H3BO3, 6 mg; CoCl2 · 6H2O, 190 mg; CuCl2 · 2H2O, 2 mg; NiCl2 · 6H2O, 24 mg; and Na2MoO4 · 2H2O, 36 mg. First dissolve FeCl2 in HCl, then dilute in water, add and dissolve the other salts. The vitamin solution contains (mg per liter of distilled water): vitamin B12, 50 mg; pantothenic acid, 50 mg; riboflavin, 50 mg; pyridoxamine-HCl, 10 mg; biotin, 20 mg; folic acid, 20 mg; nicotinic acid, 25 mg; nicotinamide, 25 mg; α-lipoic acid, 50 mg; p-aminobenzoic acid, 50 mg; and thiamine-HCl · 2H2O, 50 mg. Add potentially toxic carbon sources at low concentrations (toluene, 5 ppm; phenol, 1 mM; and Nabenzoate, 5 mM) and spike cultures again with the carbon source after its degradation. Use ascorbic acid (4 mM) to reduce the medium. Carry out enrichment in anoxically sealed serum bottles. When using rich sediments as an inoculum, deplete when necessary from readily oxidizable carbon sources by repeated incubation in medium free of carbon sources (Fries et al., 1994). Plate the enriched samples on agar media such as M-R2A medium (Fries et al., 1994) or on agar used for dilution series (Widdel and Bak, 1992). M-R2A medium contains (per liter of distilled water): yeast extract, 0.5 g; peptone, 0.5 g; casamino acids, 0.5 g; glucose, 0.5 g; soluble starch, 0.5 g; sodium pyruvate, 0.3 g; K2HPO4, 0.4 g; KH2PO4, 0.25 g; KNO3, 0.505 g; CaCl2 · 2H2O, 0.015 g; MgCl2 · 2H2O, 0.02 g; FeSO4 · 7H2O, 0.007 g; Na2SO4, 0.005 g; NH4Cl, 0.8 g; MnCl2 · 4H2O, 5 mg; H3BO3, 0.5 mg; ZnCl2, 0.5 mg; CoCl2 · 6H2O, 0.5 mg; NiSO4 · 6H2O, 0.5 mg; CuCl2 · 2H2O, 0.3 mg; NaMoO4 · 2H2O, 0.01 mg; and agar, 15 g. Adjust the pH to 7.0 before autoclaving.

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

For cultivation on poorly water-soluble compounds such as alkylbenzenes, special procedures are required (Rabus and Widdel, 1995).

Identification Since members of the genus Azoarcus are quite heterogeneous with respect to ecology and physiology, the description of the genus is complex. The phenotypic characteristics of the genus Azoarcus are as follows: Cells are straight to slightly curved rods. Cell pairs often appear slightly S-shaped, with the exception of A. buckelii cells, which occur as cocci. Cell width is 0.4– 1.5 µm and length 1.1–4.0 µm. In many species, some elongated cells (8–12 µm) occur in late-log or stationary-phase cultures on medium containing malic acid and molecular nitrogen (N2) or ammonia as nitrogen source. Cells accumulate polyhydroxybutyrate granules and are Gram negative. Cells are highly motile by means of one, rarely two polar flagella. Many species fix nitrogen and then require microaerobic conditions for growth on N2. On semisolid nitrogenfree media, microaerophilic growth can be observed as veil-like pellicles developing several mm below the surface and moving to the medium surface during growth. Colony morphology differs in plant-associated and soilborne species. Azoarcus communis, A. indigens and Azoarcus sp. BH72 grow well on complex media such as trypticase soy agar (TSA) or VM medium containing beef extract or peptone, yeast extract, and ethanol (colony diameter 2– 4 mm after 4 d); on the latter medium, they produce a yellowish, nondiffusible pigment. In contrast, the soil-borne species A. tolulyticus, A. evansii, A. buckelii, A. toluvorans and A. toluclasticus grow poorly or not at all on complex media, and some may show different pigmentation. Azoarcus anaerobius is an exception since it is strictly anaerobic and therefore does not develop colonies on aerobically incubated agar A

Fig. 2. Microcopic images of Azoarcus spp. Transmission electron (A) and phase contrast (B, C) images. A) Azoarcus sp. strain BH72 cultured in liquid VM medium for 24 h. B) Azoarcus indigens VB32T cultured on N2 in semisolid SM medium for 24 h. C) Azoarcus tolulyticus BL2 grown on nutrient plates for 48 h. Bars, 5 µm. A) and B) are from Reinhold-Hurek et al. (1993) and C) is from Song et al. (1999).

C

877

plates. Optimum temperature for growth is 28–40°C, and no growth occurs at 45°C (Fig. 2). The soil-borne species are chemoorganoheterotrophic. Bacteria have a strictly respiratory metabolism with O2 as the terminal electron acceptor, except one species (A. anaerobius which is strictly anaerobic). Alternatively, under anaerobic conditions, nitrate can be used for dissimilatory nitrate reduction. Nitrate can be also assimilated. Bacteria are oxidase positive. They grow well on salts of organic acids (such as Lmalate, succinate, fumarate and DL-lactate), ethanol and L-glutamate, but not on mono- or disaccharides except for species that are not plant-associated. These soil-borne species use few carbohydrates and a variety of aromatic substrates as sole carbon sources under denitrifying conditions. Growth factor requirement varies: some strains depend on p-aminobenzoic acid or on cobalamin. All investigated species have 16:1, and all except one cis-9 16:1 and 18:1 as major cellular fatty acids. The mol% G+C of the DNA is 62–68 (Tm). The type species is Azoarcus indigens. Since the genera Azovibrio, Azospira and Azonexus were formerly included in Azoarcus sensu lato (Reinhold-Hurek and Hurek, 2000) and are physiologically very similar, some characteristics distinguishing these genera from each other are given in Table 1. Genera harboring plant-associated diazotrophs and Thauera were also included in Table 1. The genus Thauera (Macy et al., 1993; Anders et al., 1995), which is phylogenetically closely related, is physiologically similar to soil-borne species and thus difficult to distinguish by classical tests. The validly described species of Azoarcus are A. indigens (type species) and A. communis (Reinhold-Hurek et al., 1993), A. tolulyticus (Zhou et al., 1995), A. toluclasticus and A. toluvorans (Song et al., 1999), A. evansii (Anders et al., 1995), A. anaerobius (Springer et al., 1998) and A. buckelii (Mechichi et al., 2002). These species can be distinguished from each other B

β + 0.6–0.8 Beige − − − − − − − − − + − −

β da 0.4–1.0b Yellowish to beigec − da −

+f +g +h +i +k d d −l d

Subclass Cells curved Cell width (µm) Colony color

− − − − + + + − −

β + 0.4–0.6 Pinkish translucent − − −

Azospira

− − − − + − − − +

− − +

β + 0.6–0.8 Ocher

Azonexus

d + − − + − + + +

− + −

β + 0.6–0.7 Cream

Herbaspi-rillum

− + + + d + + + +

− + −

β − 0.3–0.8 Cream

Burkholderia vietnamiensis

ND ND − ND d ND + d +

α + 0.8–1.4 Pinkish opaque d + −

Azospirillum

a

Symbols and abbreviations: +, positive for all strains; −, negative for all strains; d, 11–89% of strains are positive; and ND, not determined. Positive for soil-borne species, negative for plant-associated species. b Except for A. anaerobius and A. buckelii (1.5 µm). c Several strains do not grow on nutrient agar. d Tested for T. aromatica. e Except for T. selenatis. f ND for A. toluvorans, A. toluclasticus, A. anaerobius, A. buckelii. g Negative for A. evansii. h Negative for A. toluclasticus, d for A. tolulyticus. i Negative for A. indigens, ND for A. toluvorans, A. toluclasticus, A. anaerobius. j ND for T. mechernichensis. k Negative for A. tolulyticus, ND for A. toluvorans, A. toluclasticus, A. anaerobius, A. buckelii. l ND for A. buckelii.

Fermentative Growth on sugars Requirement for cobalamine Growth in presence of O2 on n-Butylamine 4-Hydroxybenzoate Phenylacetate Glutarate 2-Oxoglutarate n-Caproate Proprionate D-Mannose L-Proline

Azovibrio

Azoarcus

Character

ND ND ND ND ND ND ND ND +

+ + −

α − 0.7–0.9 Brown

Gluconacetobacter diazotrophicus

Thauera

ND + + +j ND d +d ND +d

−d −e −

β − 0.7–1.0 White-yellowishc

Table 1. Differential characteristics of the genera Azoarcus, Azovibrio, Azospira, Azonexus and physiologically similar diazotrophs or related bacteria of the Proteobacteria.

878 B. Rareinhold-Hurek and T. Hurek CHAPTER 3.2.18

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

by phenotypic characteristics given in Table 2. Strain BH72 is included in Table 2 and may represent a species of its own according to DNADNA-similarity studies (Reinhold-Hurek et al., 1993). It has to be noted, however, that in some cases characteristics have been tested for one strain only (see also Table 3). A more complete list of the characteristics of these species is given in Table 3. Further comments on the species are given in the text below. Azoarcus indigens is the type species and can be differentiated from the others by its requirement for p-aminobenzoic acid, growth on itaconate, and a combination of characteristics given in Table 2. Cells are thin (0.5–0.7 µm wide) and curved, and cell pairs look slightly S-shaped. Colonies are very compact and difficult to disperse; growth in liquid media is clumpy; aggregation is very strong on peptone media. Optimum temperature for growth is 40°C. The major fatty acids are 3-OH-10:0, 12:0, cis-9 16:1, cyclo-17:0, and 18:1. Azoarcus indigens was isolated from roots and stem bases of Leptochloa fusca (L.) Kunth from the Punjab of Pakistan (ReinholdHurek et al., 1993), and black sclerotia of an Ustilago-related basidiomycete from rice soil in the Punjab of Pakistan (Hurek et al., 1997b) and from rice roots (Oryza sativa) from Nepal (Engelhard et al., 2000). The type strain is strain VB32 (=LMG 9092), which has a G+C content of 62.4 mol% (Tm). Accession number of the 16S rDNA sequence is AF011345 (www.ncbi.nlm. nih.gov/entrez/query.fcgi?db=nucleotide=search =AF011345). Azoarcus anaerobius can be distinguished from others by its strictly anaerobic life style with nitrate as the only electron acceptor. Nitrate is quantitatively reduced to N2 gas, nitrite accumulates intermediately, and nitrous oxide (N2O) is not detected. Additional differences are given in Table 2. Optimum growth temperature is 28°C. Enhanced salt concentration impairs growth. Cells are superoxidedismutase-positive. Sulfate, thiosulfate, sulfite, sulfur, trimethyl-amine N-oxide, dimethyl sulfoxide (DMSO), Fe(OH)3, K3[Fe(CN)6], or fumarate are not reduced, and oxygen is not reduced, not even at low pressures. Characteristics in addition to those given in Table 3 are: Sole carbon sources for growth are propanol, valerate, pyruvate, cyclohexane carboxylate, phenol, resorcinol, p-cresol; no growth with L-malate, formate, 5-oxocaproate, pimelate, catechol, hydroquinone, 2-hydroxybenzoate, ocresol, and m-cresol. No autotrophic growth with hydrogen or thiosulfate. The pH range is 6.5–8.2 and optimum pH 7.2. Cells were isolated from sewage sludge. The species consists of only one strain, which is the type strain LuFRes1 (= DSM 12081). It has a G+C content of 66.5 ±

879

0.5 mol% (Tm). Accession number of the 16S rDNA sequence is Y14701 (www.ncbi.nlm.nih. gov/entrez/query.fcgi?db=nucleotide=search= Y14701). Azoarcus buckelii can be distinguished from others by its cell shape and a combination of characteristics given in Table 2. Cells are coccoid of irregular size (1.5 ± 0.5 µm). Optimum temperature for growth is 28°C with a range from 4 to 40°C. It requires vitamin B12 for growth. Also nitrite can be used as terminal electron acceptor. Under oxic conditions, methanol and pyruvate are utilized in addition to carbon sources listed in Table 3. No growth occurs on formate, 2-aminobenzoate, catechol, pimelate, hexadecane, o-cresol, p-cresol, cyclohexane carboxylate, and phenol. Under denitrifying conditions, they utilize benzoate, 3-hydroxybenzoate, 4-hydroxybenzoate, 2aminobenzoate, o-phtalate, protocatechuate, phenol, o-cresol, p-cresol, cyclohexane carboxylate, 4-hydroxyphenylacetate, heptanoate, 2hydroxybenzaldehyde, 3-hydroxybenzaldehyde, 4-hydroxybenzaldehyde, 2-hydroxybenzoate, 3hydroxybenzoate, 4-hydroxybenzoate, 2,4dimethylphenol, 4-hydroxy-3-methylbenzoate, or for slow growth, 3-methylbenzoate, 2fluorobenzoate, gentisate, phenylacetate, phenylpropionate, L-tyrosine, L-phenylalanine, and indole 3-acetate; no growth occurs on m-cresol, 3-aminobenzoate, 4-aminobenzoate, 2methylbenzoate, 4-methylbenzoate, catechol, resorcinol, hydrochinone, cinnamate, p-anisate, L-tryptophan, indole, ethylbenzene, benzene, 3chlorobenzoate, adipate, pimelate, D-glucose, D-fructose, D-ribose, lactose, cyclohexanol, cyclohexanone, 1,3-cyclohexanedione, D,Lmandelate, D,L-4-hydroxymandelate, and 2hydroxybenzyl alcohol. The species consists of only one strain, which is the type strain U120 (= DSMZ 14744). It has a G+C content of 66 mol%. Accession number of the 16S rDNA sequence is AJ315676 (www.ncbi.nlm.nih.gov/ entrez/query.fcgi?db=nucleotide=search= AJ315676). Azoarcus communis can be distinguished from others by its cell width (0.8–1.0 µm) in combination with growth on D-mandelate, by growth on citrate, and a combination of characteristics given in Table 2. Cells are only slightly curved and plump. Optimum temperature for growth is 37°C. Some strains still grow well in 2% NaCl. The major fatty acids are 3-OH-10:0, 14:0, cis-9 16:1, 16:0, and 18:1. This species was isolated from roots of L. fusca (L.) Kunth from the Punjab of Pakistan (SWub3T; Reinhold-Hurek et al., 1993; Engelhard et al., 2000) and from refinery oily sludge in France (Laguerre et al., 1987). The species affiliation of strain LU1 isolated from a compost biofilter in Canada (Juteau et al., 1999)

0.8–1.0 + + + − + + + d + + − − − − − − − + + −

+ + + + − − − − − − − + + + − −

A. communis

0.5–0.7 +a + + +

A. indigens

d d d − d + + + + + + − − − − +

0.8–1 + d + −

0.6–0.8 + + + − + + − + + + − − − − − − − − − −

A. tolulyticus

A. sp. BH72

− + ± − − ± + + − − + ND ND ND ND +

0.8–1.0 + + ND −

A. toluvorans

− − d − − − + + d d + ND ND ND ND +

0.6–0.8 + − ND −

A. toluclasticus

+ d + + − + + d d + − − + − − −

0.4–0.8 + + − −

A. evansii

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

1.5 − − − −

A. anaerobius

Symbols and abbreviations: +, positive for all strains; −, negative for all strains; ±, weak reaction for all strains; d, 11–89% of strains are positive; and ND, not determined.

Cell width (µm) O2 terminal electron acceptor Catalase Nitrogen fixation Requirement for p-aminobenzoic acid Sole carbon sources for growth 3-Hydroxybenzoate Phenylacetate L-Phenylalanine D-Tartrate n-Caproate Isovalerate L-Proline, D(+)-galactose, sucrose Maltose D(+)-Glucose D(+)-Fructose D(−)-Ribose, adipate Itaconate p-Aminobenzoate D-Mandelate Citrate L(+)-Arabinose, D(+)-xylose

Characteristic

Table 2. Characteristics differentiating species of the genus Azoarcus from each other.

+ ± + ND − ND ND ND − − ND ND ND ND − ND

1.5 + + − −

A. buckelii

880 B. Rareinhold-Hurek and T. Hurek CHAPTER 3.2.18

Colony surface Growth at 40°C Requirement for p-aminobenzoic acid Requirement for cobalamin Nitrogen fixation Oxidation/fermentation of glucose Catalase Sole carbon sources for growthg n-Butylamine 3-Hydroxybenzoate Benzylamine Phenylacetate Benzoate L-Phenylalanine

Colony color (CR)c

Cell width (µm) Cell length (µm) Elongated cells in stationary cultures Colony diameter (mm, VM/CR)c Colony color (VM)c

Characteristic

2–4

Translucent yellowish Whiteish pink pink center Smooth + −

d

+ −

+

+ + v + d +

Opaque yellowish Whiteish pink white margin Rough + +

−

+ −

+

+ + d + d +

0.8–1.0 1.5–3.0 r

Azoarcus communis (n = 3)

2–3

0.5–0.7 2.0–4.0 r

Azoarcus indigens (n = 5)

+ + + + + −

+

+ −

−

Shining + −

Translucent yellowish Orange red

2–3/1

0.6–0.8 1.5–4.0 r

Azoarcus sp. BH72 (n = 1)

+b d +b d + d

D

+ ND

−

Shining −b −

Opaque yellowishb Orange redb

ND − ND + + ±

+

ND ND

−

ND − −

ND

ND

ND

0.8–1.0 1.4–2.8 ND

0.8–1b 1.4–2.8 − 1–1.5b

Azoarcus toluvorans (n = 2)

Azoarcus tolulyticusb (n = 9)

ND − ND − + d

−

ND ND

−

ND − −

ND

ND

ND

0.6–0.8 1.7–4.0 ND

Azoarcus toluclasticus (n = 5)

Table 3. Characteristics of species of the genera Azoarcus, Azovibrio, Azospira and Azonexus.a

+b +b +b d + +

+

+ +d

−

Shining −b −

Translucent beigeb Orange redb

0.2–0.7b

0.4–0.8 1.5–3.0 −

Azoarcus evansiib (n = 2)

ND + ND ± + +

+

− ND

+

ND + −

ND

ND

Shining d −

− − −

ND + ND + + +

−

− −

−

− − − − − −

+

+ −

−

Opaque beige Orange red

− −

1.5–2/1

0.6–0.8 1.5–3.5 r

Azovibrio restrictus (n = 5)

0

1.5 2.7–3.3 ND

1.5 ± 0.5 1.5 ± 0.5 − ND

Azoarcus anaerobius (n = 1)

Azoarcus buckelii (n = 1)

− − − − − −

ND

+ −

+

ND ND −

Negligible

Negligible

Negligible

0.6–0.8 1.5–3.5 ND

Azovibrio sp. BS20-3 (n = 1)

− − − − − −

+

+ −

d

Shining + −

Translucent pink-salmon Translucent orange

1–2

0.4–0.6 1.1–2.5 r

Azospira oryzae (n = 12)

− − − − − −

+

+ −

+

Shining + −

Opaque ocher Dark red

1–2/0.7

0.6–0.8 1.5–4.0 +

Azonexus fungiphilus (n = 5)

CHAPTER 3.2.18 The Genera Azoarcus, Azovibrio, Azospira and Azonexus 881

D(+) Malate L-Alanine 2-Oxoglutarate D-Tartrate n-Caproate Propionate Isovalerate L-Proline Malonate Maltose D(+)-Glucose D(+)-Fructose D(+)-Galactose, sucrose D(−)-Ribose Maltotriose, palatinose, D(+)-melezitose DL-Glycerate Toluene (denitrifying) Protocatechuate, L-mandelate 4-Hydroxybenzoate m-Coumarate Tryptamine Gentisate L(+)-Tartrate meso-Tartrate, betaineb p-Aminobenzoate Itaconate Glutarate D-Mandelate

Characteristic

Table 3. Continued

− − − − +

− −

− − +

+ + v + − − − − + +

− −

d − +

+ d + + − − + + − +

+f −b −b −b − − −b −b +b −b

+b + −b

+ +

+b +b −b − d d + +b − + + + +

Azoarcus tolulyticusb (n = 9)

ND d ND +f ND ND ND − ND ND ND ND ND

+f ND ND ND − ND ND ND ND ND

+ ND

ND ND ND − − − − + ND + d d +

Azoarcus toluclasticus (n = 5)

ND + ND

+ ND

ND ND ND − − − ± + ND + − − +

Azoarcus toluvorans (n = 2)

± ND ND − ND ND ND ND + ND

ND ND ND

+b ND −b − −b −b + +b +b + −b +b −b

ND ND

ND ND ND ND +f ND ND ND ND ND − − ND

Azoarcus buckelii (n = 1)

−b +b

−b +b +b + −b + +b + − d d +b +b

Azoarcus evansiib (n = 2)

+f ND ND ND ND ND ND ND ND ND

ND ND ND

ND ND

− ND ND ND ND + + ND ND ND − − ND

Azoarcus anaerobius (n = 1)

−b − − − − − − − − −

− − − − − − − − − −

− − −

− −

− − − − −

− + − − − + + − − − − − −

Azovibrio sp. BS20-3 (n = 1)

D D − − − + − − − − − − −

Azovibrio restrictus (n = 5)

− − − − − − − − − −

− − −

− −

− − + + + + + − − − − − −

Azospira oryzae (n = 12)

− − − − − − − − − −

− − −

− −

− − + − − − − + d − − − −

Azonexus fungiphilus (n = 5)

B. Rareinhold-Hurek and T. Hurek

+ + + + − − − − + −

+ + + + + + + − − − − − −

+ + + d + + + − − − − − −

+ d + + − + − − − − − − −

Azoarcus sp. BH72 (n = 1)

Azoarcus communis (n = 3)

Azoarcus indigens (n = 5)

882 CHAPTER 3.2.18

+ d d + v + + + − − − −

−

− − − − + + + + − − − −

−

−

− − − + + + + + − − − −

Azoarcus sp. BH72 (n = 1)

+

− − +b +b +b +b + + + − − +

Azoarcus tolulyticusb (n = 9)

d

ND ND ND ND + ND ND + + + − +

Azoarcus toluvorans (n = 2)

+

ND ND ND ND + ND ND d + + − +

Azoarcus toluclasticus (n = 5)

−

− − − +b + +b + + − − − −

Azoarcus evansiib (n = 2)

ND

− +h ND ND ± ND + ND ND ND ND ND

Azoarcus buckelii (n = 1)

ND

ND ND ND ND ND ND − + ND ND + −

Azoarcus anaerobius (n = 1)

−

− − − d + − + + − − − −

Azovibrio restrictus (n = 5)

−

− − − + − + + + − − − −

Azovibrio sp. BS20-3 (n = 1)

−

− − − v + + + + − − − −

Azospira oryzae (n = 12)

−

− − − − + + + + − − − −

Azonexus fungiphilus (n = 5)

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Symbols and abbreviations: +, positive for all strains; −, negative for all strains; ±, weak reaction for all strains; d, 11–89% of strains are positive; r, rarely; v, variable result; and ND, not determined. a All strains have the following features: cells are straight to curved rods except A. buckelii; oxidase positive; no growth in the presence of 5% NaCl and no growth-rate increase when NaCl is added to medium (ND for A. toluvorans, A. toluclasticus, A. anaerobius and A. buckelii); denitrification (ND for Azonexus sp.); no spore formation; no starch hydrolysis (ND for A. evansii, A. buckelii and A. anaerobius). b Marked characters were determined for A. tolulyticus strain Td-1 or A. evansii strain KB740T (Reinhold-Hurek and Hurek, 2000). c Growth on VM ethanol agar (VM) or Congo red agar (CR) at 37°C for 4 d. d Not tested for strain pF6. e All strains grow on DL-lactate, succinate, acetate, L-glutamate, butyrate (ND for A. buckelii), ethanol (ND for A. toluvorans and A. toluclasticus; no substrate but L-malate tested for A. evansii pF6); no growth on (ND for A. buckelii) D(+)-mannose, maltitol, N-acetyl-D-glucosamine, D-gluconate, and caproate; no growth onb (ND for A. toluclasticus, A. toluvorans, A. anaerobius and A. buckelii) D(+)-trehalose, L(+)-sorbose, ∀-D(+)melibiose, D(+)-raffinose, lactulose, 1-0-methyl-β-galactopyranoside, 1-0-methyl-∀-galactopyranoside, D(+) cellobiose, β-gentiobiose, 1-0-methyl-β-D-glucopyranoside, ∀-L-rhamnose, ∀-L-fucose, D(+)-arabitol, L(−)-arabitol, xylitol, dulcitol, D-tagatose, myo-inositol, D-sorbitol, adonitol, hydroxyquinoline-β-glucuronide, D-lyxose, erythritol, 1-0-methyl-∀-D-glucopyranoside, 3-0-methyl-glucopyranose, D-saccharate, mucate, D-glucuronate, Dgalacturonate, 2-keto-D-gluconate, 5-keto-D-gluconate, 3-phenylpropionate, caprylate, 4-aminobutyrate, DL-∀-amino-n-valerate, trigonelline, putrescine, ethanolamine, and D-glucosamine. f Under denitrifying conditions. g Tested with O2 as terminal electron acceptor. h Only glycerol tested. From Reinhold-Hurek et al. (1993), Anders et al. (1995), Hurek and Reinhold-Hurek (1995a), Zhou et al. (1995), Hurek et al. (1997), Rhee et al. (1997), Springer et al. (1998), Song et al. (1999), Engelhard et al. (2000), Reinhold-Hurek and Hurek (2000), and Mechichi et al. (2002).

Citrate Glycerol, D-alanine (−)-Quinate Isobutyrate L-Aspartate 3-Hydroxybutyrate L-Malate Fumarate Adipate L-Arginine L-Tyrosine L(+) Arabinose, D(+)-xylose ∀-Lactose

Characteristic

Azoarcus communis (n = 3)

Azoarcus indigens (n = 5)

CHAPTER 3.2.18 883

884

B. Rareinhold-Hurek and T. Hurek

is currently not clear. The type strain is strain SWub3 (= LMG 9095), which has a G+C content of 62.4 mol% (Tm). Accession number of the 16S rDNA sequence is AF011343 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=AF011343). Azoarcus evansii can be distinguished from other species by a combination of characters given in Table 2. Cells are rods with rounded ends and motile by means of a subpolarly inserted flagellum. Yeast extract (0.1% [w/v]) inhibits growth. Optimum growth temperature is 35–37°C, and optimum pH is 7.8. During denitrifying growth with aromatic compounds, nitrite is an intermediate and is reduced mainly to N2O (strain KB740T). Characteristics in addition to those listed in Table 3 are: Under anaerobic conditions, strain KB740T uses benzoate, phenylacetate, phenylglyoxylate, 3- and 4hydroxybenzoate, 2-aminobenzoate, 4-hydroxyphenylacetate, L-phenylalanine, p-cresol, 2fluorobenzoate, benzaldehyde, benzyl alcohol, indolylacetate, o-phtalate, adipate, pimelate, cyclohexane carboxylate, succinate, fumarate, Lmalate, acetate, acetone, D-fructose, D-maltose, and slow growth occurs on glutarate, D-glucose, but not on toluene, phenol, 2-hydroxybenzoate, protocatechuate, o- and m-cresol, indole, ethanol, D-ribose, or D-lactose. Pyridine is used by strain pF6 under aerobic and anaerobic conditions. The major cellular fatty acids are cis-9 16:1, 16:0, 12:0, 3-OH 10:0, and cis-9, 11 18:1. The major quinone is ubiquinone 8. The species description is based on one strain isolated from creek sediment in the United States (Braun and Gibson, 1984); however, a second strain, pF6, sharing 100% 16S rDNA sequence identity was isolated on pyridine from industrial wastewater in Korea (Rhee et al., 1997). The type strain is strain KB740 (= DSM6898), which has a G+C content of 67.5 mol%. Accession number of the 16S rDNA sequence is X77679 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=X77679). Azoarcus toluclasticus can be differentiated from the other species by the lack of catalase activity despite aerobic growth and by a combination of characters given in Table 2. Cells are short motile rods. Optimum temperature for growth is 30°C. Under denitrifying conditions, they can grow on acetate, benzoate, pyruvate, succinate, D-xylose, L-arabinose, D-ribose, Dgalactose, sucrose, lactose, maltose, adipate, lactate, mannitol, L-aspartate, L-proline or Larginine. All strains except strain MF23 can use toluene as a growth substrate and strains MF58 and MF63T can also grow on phenol under denitrifying conditions. The strains grow on brainheart infusion, nutrient and trypticase soy agar (TSA), except for strain MF63T, which does not

CHAPTER 3.2.18

grow on nutrient agar. The predominant cellular fatty acids are 16:0 and 16:1ω7c. DNA-DNA hybridizations show intermediate similarities (47–55%) among genomovar I (strains MF7 and MF23) and genomovar II (strains MF58, MF63T and MF441). They were isolated from aquifer sediments in the United States. The type strain is strain MF63 (= ATCC 700605), which has a G+C content of 67.3 mol% (HPLC). Accession number of the 16S rDNA sequence is AF123077 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=AF123077). Azoarcus tolulyticus can be differentiated from the other species by a combination of characters given in Table 2. Cells are short motile rods, which are slightly elongated (to 2.8 µm) when grown on M-R2A agar. Optimum temperature for growth is 30°C, and growth still occurs at 37°C. Under denitrifying conditions they can utilize acetate, adipate, L-arginine, L-arabinose, L-aspartate, benzoate, fumarate, D-galactose, Dglucose, lactate, lactose, maltose, mannitol, Lproline, pyruvate, D-ribose, succinate, sucrose, toluene, D-xylose, or 4-hydroxybenzoate. The strains grow on brain-heart infusion agar, but not or only poorly on nutrient and TSA. The predominant cellular fatty acids are 16:0 and 16: 1ω7c. Strains Td-19 and Td-3 (Zhou et al., 1995) have been removed from the species owing to low DNA-DNA similarity values (Song et al., 1999). The source of strains is aquifer sediments and petroleum-contaminated soils in the United States (Fries et al., 1994; Fries et al., 1997; Chee-Sanford et al., 1996). The type strain is strain Tol-4 (= ATCC 51758), which has a G+C content of 66.9 mol% (HPLC). Accession number of the 16S rDNA sequence is L33694 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=L33694). Azoarcus toluvorans can be differentiated from the other species by a combination of characters given in Table 2. Optimum temperature for growth is 30°C. Grow on brain-heart infusion, nutrient and TSA. Under denitrifying conditions, they can grow on acetate, benzoate, butyrate, fumarate, phenylacetate, pyruvate, succinate, toluene, D-xylose, L-arabinose, D-ribose, Dgalactose, sucrose, maltose, adipate, lactate, mannitol, L-aspartate, L-proline, L-phenylalanine, L-arginine, 4-hydroxybenzoate or phenol, and under aerobic conditions, on benzene or ethylbenzene. They were isolated from soil from an industrial area in Brazil (Td17) or noncontaminated organic soil in the United States (Td21T; Fries et al., 1994). The type strain is strain Td21 (= ATCC 700604), which has a G+C content of 67.8 mol% (HPLC). Accession number of the 16S rDNA sequence is L33692 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=L33692).

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Azoarcus sp. strain BH72 can be differentiated from the other species by its lack of growth on L-phenylalanine and a combination of characteristics given in Table 2. Cells are long, thin (0.6– 0.8 µm wide) and slightly curved, elongated (8–12 µm) cells occurring in late-log or stationary phase culture on N-free or ammoniumsupplemented SM-medium. Optimum temperature for growth is 40°C. The major fatty acids are16:0, cis-9 16:1, 18:1, 18:1, and 14:0. It was isolated from the root interior of Leptochloa fusca (L.) Kunth from the Punjab of Pakistan (Reinhold et al., 1986). The G+C content is 67.6 mol% (Tm). Accession number of the 16S rDNA sequence is AF011344 (www.ncbi.nlm. nih.gov/entrez/query.fcgi?db=nucleotide=search =AF011344).

Preservation Although organic acids are typical carbon sources for many strains, medium containing organic acids becomes alkaline in the stationary phase. Thus, subculturing on organic acids should be avoided and VM ethanol medium should be used instead. Soil-borne strains may also be subcultured on M-R2A medium. Preservation by lyophilization is possible. Long-term preservation is achieved by storage in liquid nitrogen with 5% DMSO as a cryoprotectant.

Physiology The optimum growth temperature is 30°C or 37– 40°C, which is lower for A. anaerobius (28°C). Optimum pH is neutral, and the bacteria are not halophilic. All Azoarcus species have a strictly respiratory type of metabolism, and no fermentative abilities have been observed. All species except one, A. anaerobius, use oxygen as the terminal electron acceptor. In the original description of Azoarcus spp., standard procedures had failed to reveal the capacity for denitrification (Reinhold-Hurek et al., 1993). In a closer examination, the capacity to use nitrate as the terminal electron acceptor was demonstrated for these plant-associated strains (Hurek and ReinholdHurek, 1995a), a feature which they share with all other species (Anders et al., 1995; Zhou et al., 1995; Springer et al., 1998; Song et al., 1999). Most strains belonging to A. tolulyticus, A. evansii, A. toluclasticus, A. toluvorans and A. anaerobius were enriched or isolated anaerobically on nitrate, whereas the plant-associated species were enriched under conditions of nitrogen fixation. In the original description of the genus Azoarcus, the capacity to fix nitrogen is listed as a genus character (Reinhold-Hurek et al., 1993). Nitrogen fixation (Fries et al., 1994; Zhou et al.,

885

1995) or the occurrence of a nitrogenase gene nifH (Hurek et al., 1997a) has also been demonstrated for A. tolulyticus. However, owing to the addition of new species, this cannot be generalized for Azoarcus anymore. Nitrogen fixation was not detected in A. anaerobius (Springer et al., 1998) or A. buckelii (Mechichi et al., 2002). In the absence of physiological data (Anders et al., 1995), A. evansii was thought unlikely to be diazotrophic since no polymerase chain reaction (PCR)-amplifiable nifH fragment (T. Hurek and B. Reinhold-Hurek, unpublished observations) or hybridization with a nifH probe (Hurek et al., 1997a) occurs. However, recently this species was reported to fix nitrogen (Mechichi et al., 2002). For most other isolates enriched under denitrifying conditions, this character was not tested at all. In all diazotrophic strains (A. indigens, A. communis, A. tolulyticus and Azoarcus sp. strain BH72), nitrogenase activity occurs only under microaerobic conditions, probably due to the lack of efficient oxygen protection mechanisms. It was shown that nitrogen fixation is more tolerant to oxygen in strain BH72 than in Azospirillum spp., reaching steady states in an oxygen-controlled chemostat until dissolved O2 exceeds 25 µM (Hurek et al., 1987). Similarly, the expression of nitrogenase genes is transcriptionally regulated in response to oxygen (fully repressed at 4% oxygen in the headspace) or combined nitrogen (repressed by 0.5 mM ammonium or nitrate; Egener et al., 1999). This strain shows augmented rates and efficiency of nitrogen fixation, called “hyperinduction,” in empirically optimized batch cultures when shifting down to extremely low oxygen concentrations (30 nM; Hurek et al., 1994a). In the course of hyperinduction, novel intracytoplasmic membrane stacks are formed which might participate in efficient nitrogenase activity, since the iron protein of nitrogenase is mainly associated with these membranes (Hurek and Reinhold-Hurek, 1995a). The formation of these so-called “diazosomes” is most abundant and reproducible in coculture of strain BH72 and endophytic Acremonium alternatum, a mitosporic deuteromycete related to the Hypocreales within the euascomycetes and an isolate from Kallar grass (Hurek et al., 1995b; Hurek and Reinhold-Hurek, 1998). As in many Proteobacteria, nitrogenase genes are organized in an nifHDK operon (Egener et al., 2001). Phylogenetically, the nitrogenase in the genus Azoarcus follows either the organismic phylogenetic tree or appears to have been obtained by lateral gene transfer, depending on the species. In A. indigens, A. communis and Azoarcus sp. strain BH72, a fragment of the iron protein of nitrogenase encoded by the nifH gene is most closely related to nitrogenases occurring in diazotrophs of the γ-subclass of Proteobacte-

886

B. Rareinhold-Hurek and T. Hurek

ria, while in A. tolulyticus, it is located within a clade of Alphaproteobacterial nitrogenases (Bradyrhizobium and Azospirillum; Hurek et al., 1997a). In general, carbohydrates are not the preferred carbon sources of Azoarcus spp. None of the plant-associated species is able to utilize any out of 50 mono- and disaccharides or sugar alcohols tested (Reinhold-Hurek et al., 1993). Azozrcus anaerobius shows no growth on common carbohydrates such as D(+)-glucose or D(–)fructose (Springer et al., 1998). In contrast, among the soil-borne species, all strains tested so far are able to utilize at least some carbohydrates (Table 1). All strains grow well on organic acids and a few amino acids (Table 1). The carbon sources listed here (except for A. anaerobius) were tested with O2 as the terminal electron acceptor. For some strains, tests were also carried out under anaerobic conditions with nitrate as electron acceptor. For the majority of carbon sources, results were identical; however, for several carbon sources, strain-dependent differences were found (Song et al., 1999). In contrast to plant-associated species (Reinhold-Hurek et al., 1993; Hurek and Reinhold-Hurek, 1995a), most soil-borne species grow on aromatic compounds (such as toluene or phenol) under denitrifying conditions (Zhou et al., 1995; Song et al., 1999), benzoate (Anders et al., 1995), or resorcinol (Springer et al., 1998). Owing to the anaerobic degradation of aromatic compounds, this bacterial group receives particular attention for their biodegradation and biotransformation abilities. While aerobic metabolism is characterized by the extensive use of molecular oxygen (which is essential for the hydroxylation and cleavage of the ring structures), different strategies are necessary for the anaerobic degradation and are currently under study. Toluene, which can be decomposed anaerobically by three species, is activated by the addition to fumarate to form benzylsuccinate (Beller and Spormann, 1997) catalyzed by benzylsuccinate synthase via a glycyl radical (Krieger et al., 2001). The aerobic metabolism of benzoate is unusual in A. evansii KB740T. None of the known pathways, including the conversion of benzoate to catechol (1,2-dihydroxybenzoene) or protocatechuate (3,4-dihydroxybenzoate), appears to operate in this species (Mohamed et al., 2001). The first step is the activation of benzoate to benzoyl-CoA by a benzoate-CoA ligase, and the second step involves the hydroxylation of benzoyl-CoA by a novel benzoyl-CoA oxygenase (Mohamed et al., 2001). The first step of phenylacetate degradation is catalyzed by two different phenylacetate-CoA ligases under aerobic and anaerobic conditions, respectively (Mohamed, 2000). Strain PF6 degrades pyridine

CHAPTER 3.2.18

aerobically and anaerobically with nitrate as electron acceptor (Rhee et al., 1997). BenzoylCoA is also a central intermediate in the anaerobic degradation of aromatic compounds (Harwood et al., 1998). In A. evansii KB740, nucleotide sequence analysis of the gene cluster including a gene for benzoate-CoA ligase indicates that the degradation of benzoate is probably similar to the benzoate-CoA pathway in Thauera aromatica (Harwood et al., 1998). For further reading on anaerobic oxidation of aromatic compounds, the review of Boll et al. (2002) is recommended. Resistance of Azoarus spp. to antibiotics has not been extensively tested. Azoarcus indigens, A. communis and strain BH72 are not resistant to ampicillin, kanamycin, streptomycin, spectinomycin or tetracycline (B. Reinhold-Hurek and T. Hurek, unpublished observation).

Genetics For the three plant-associated species of Azoarcus, DNA reassociation experiments led to the estimation of a genome size from 4.5 to 5 Mb. The genome of Azoarcus sp. strain BH72 is currently sequenced (Hurek and Reinhold-Hurek, 2003), with an estimated genome size of 4.4 Mb. Nothing is known about bacteriophages of the genus Azoarcus. Also the plasmid content is not well studied. In strain BH72, plasmids could not be detected in Eckhardt gels or by pulse field gel electrophoresis (B. Reinhold-Hurek and T. Hurek, unpublished observations). Several strains of Azoarcus spp. have been shown to be transformable. Broad host range plasmids based on RK2 such as pRK290 or pAFR3 can be transferred by triparental mating and are stably replicated in strain BH72 (Egener et al., 2001). Transformation can also be achieved by electroporation (Hurek et al., 1995b), and mutagenesis by allelic exchange (Hurek et al., 1995b) or transposon insertion (Dörr et al., 1998) is established.

Ecology As already stated in the section on Habitat, the two phylogenetic clusters of Azoarcus strains appear to differ substantially in their ecology: the plant-associated and the soil-borne species. Among the plant-associated species (A. indigens, A. communis and strain BH72), most strains (except strain S2 from sludge [Laguerre et al., 1987] or strain LU1 from a compost biofilter [Juteau et al., 1999], both related to A. communis [Fig. 1]) appear to be strictly plant-associated and have never been isolated from root-free soil (Reinhold-Hurek and Hurek, 1998b). In contrast, no members of soil-borne species have been isolated from plant material as yet.

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Plant-associated Azoarcus spp. have been up to now isolated from a limited number of samples, such as Kallar grass (Leptochloa fusca L. Kunth) from the Punjab of Pakistan (Reinhold et al., 1986; Reinhold-Hurek et al., 1993), rice roots (Oryza spp.) from Nepal (Engelhard et al., 2000), or resting stages of plant-associated fungi (Hurek et al., 1997b). However they may be more widely distributed than assumed; in molecular-ecological studies on root material or fungal spores, Azoarcus 16S rDNA genes (Hurek and Reinhold-Hurek, 1995a) or nifH genes (Ueda et al., 1995; Engelhard et al., 2000; Hurek et al., 2002) have been retrieved which did not correspond to genes of cultivated strains or species. Interestingly, the sequences retrieved up to now from plant material did not cluster with genes from soil-borne species, confirming that the latter do not appear to be plant associated. Since bacteria corresponding to the detected sequences could not be isolated from the same plant samples, they may occur in a state in which they are difficult to cultivate and thus overlooked by classical microbiological techniques. This is in agreement with the observation that in an inoculation experiment of Kallar grass with strain BH72, already two months after inoculation, attempts to reisolate the bacteria failed; however, nitrogenase gene (nifH) mRNA of this species was present at high levels in plant roots, suggesting high metabolic activity (Hurek et al., 2002). Using nif – and wild type BH72, it was demonstrated by N-balance and 15N2/14N2 isotope studies that Azoarcus sp. can supply fixed nitrogen to its host plant Kallar grass (Hurek et al., 2002), an important observation with respect to the ecological role and possible applications of these bacteria. Since plant-associated Azoarcus spp. cannot be isolated from root-free soil, the question arises how do they newly infect plants in nature. Survival in the absence of plants may be achieved in association with fungal resting stages in the soil (Hurek). Black sclerotia of a basidiomycete (a relative of Ustilago hordei) collected from a Kallar grass site in Pakistan harbored culturable strains of Azoarcus indigens and of the related genera Azovibrio spp., Azospira oryzae and Azonexus fungiphilus (Hurek). This suggests that plant-associated fungi might serve as shuttle vectors for plant-associated bacterial strains. Indeed, when these sclerotia were used to inoculate surface-sterilized rice seedlings in gnotobiotic culture, diazotrophs could be isolated from roots after 2–3 weeks of incubation. The abundant diazotrophs were identified as Azospira oryzae (B. Reinhold-Hurek, unpublished results). Mechanisms of plant-microbe-interactions are best studied for Azoarcus sp. strain BH72, orig-

887

inating from the pioneer plant Kallar grass from Pakistan (Reinhold et al., 1986). In gnotobiotic cultures in the laboratory, these bacteria have a wider host range. They can invade rice roots and to a lesser extent stems (Hurek et al., 1994b), where they mainly colonize the root cortex tissue intercellularly and rarely the stele including xylem cells (Hurek et al., 1994b). Despite a high density of colonization of the root interior, the bacteria do not cause symptoms of plant disease (Hurek et al., 1994b) and thus have an endophytic and not a pathogenic lifestyle. Unlike rhizobia, they do not form an endosymbiosis in living plant cells (Reinhold-Hurek and Hurek, 1998a). Nevertheless they show endophytic nitrogen fixation, expressing nitrogenase genes in the apoplast of aerenchymatic air spaces of flooded rice seedlings (Egener et al., 1999) or field-grown Kallar grass plants (Hurek et al., 1997a). Soil-borne species occur in a wide range of environments as stated in the section on Habitat. In most cases they were readily isolated under denitrifying conditions from nonmarine habitats such as contaminated or noncontaminated soils or sediments. Since many strains can degrade a wide range of aromatic compounds—which are the second most abundant class of organic compounds in nature—they can be expected to play an important role in carbon degradation in the biosphere. Their occurrence in anoxic sediments or sewage sludge indicates that their lifestyle in situ might be anaerobic (on nitrate as terminal electron acceptor), in contrast to the plantassociated strains that might be fixing nitrogen microaerobically. Besides the numerous isolates, there are also strains which have as yet escaped cultivation. One example is a deeply-branching clade of Azoarcus sp. found in high numbers in nitrifying-denitrifying activated sludge of an industrial wastewater treatment plant (Juretschko et al., 2002).

The Genera Azovibrio, Azospira and Azonexus Introduction Bacteria of these genera were first described to belong to the genus Azoarcus sensu lato (Reinhold-Hurek et al., 1993; Hurek et al., 1997b), to group C (Azovibrio), group D (Azospira) and group E (Azonexus). The availability of new isolates recently allowed a reassessment of the genus by a polyphasic taxonomic approach, leading to the proposal of three new genera. Since they are relatively similar in physiology and ecology, they are treated together.

888

B. Rareinhold-Hurek and T. Hurek

Recently, perchlorate-reducing bacteria were described as a new genus (Dechlorosoma suillum; Achenbach et al., 2001); however, a polyphasic taxonomic analysis by us (Tan and Reinhold-Hurek, 2003) demonstrated that they are members of Azospira oryzae. Therefore, these bacteria are also included in this chapter.

Phylogeny Analysis of almost complete 16S rDNA sequences placed all three genera into the Betaproteobacteria. Phylogenetic analyses using different algorithms showed that Azoarcus sensu lato was not monophyletic, but Azoarcus sensu stricto was divided from the other groups by the Thauera clade (Reinhold-Hurek and Hurek, 2000; see Fig. 1). The three new genera clearly represent different lineages, despite their exact phylogenetic relationships within the Rhodocyclus subgroup not being well resolved. Their phylogenetic distances from each other are similar to that of the Thauera/Azoarcus group or Duganella (formerly Zoogloea; Fig. 1). Other related genera are Dechloromonas, Propionivibrio and Rhodocyclus, and none are typically plantassociated bacteria.

Taxonomy The three groups are divided from each other and to Azoarcus by a polyphasic taxonomic analysis including 16S rDNA sequence analysis, DNA-DNA hybridization, protein profiles, fatty acid analysis, and nutritional profiles. Each genus consists of only one validly described species: Azovibrio restrictus, Azospira oryzae and Azonexus fungiphilus (Reinhold-Hurek and Hurek, 2000). The members of each species are quite homogeneous in protein patterns and substrate utilization, except for Azovibrio sp. strain BS203. It shares only 97% 16S rDNA sequence identity with Azovibrio restrictus S5b2T, which is strictly microaerobic and might thus deserve the rank of a separate species in future. Azovibrio now harbors new members recently described as a misnamed genus “Dechlorosoma suillum” (Achenbach et al., 2001). The type strain shares almost 100% of 16S rDNA sequence identity (1433 out of 1435 bp) with Azospira oryzae 6a3, is very similar in sodium dodecylsulfate (SDS)-soluble protein patterns and carbon source profiles, and shows approx. 90% binding in DNA-DNA-hybridization studies; thus the type strain of Dechlorosoma suillum and Azospira oryzae have to be regarded as the same species (Tan and Reinhold-Hurek, 2003).

CHAPTER 3.2.18

Habitat and Ecology Bacteria of all three genera occur in terrestrial habitats. Azovibrio and Azospira strains have typically been isolated from surface-sterilized plant roots and are thus considered to be diazotrophic endophytes of grasses. Members of both genera were found in roots of Kallar grass (Leptochloa fusca (L.) Kunth) in the Punjab of Pakistan (Reinhold-Hurek et al., 1993) and in roots of wild rice and cultured rice Oryza sativa in Nepal (Engelhard et al., 2000). Azospira oryzae also was found in roots of cultured rice in Italy (Engelhard et al., 2000) and wild rice in the Philippines (Hurek et al., 2000; Reinhold-Hurek and Hurek, 2000). Some strains of both genera and Azonexus fungiphilus were additionally found in another microniche, the resting stages (sclerotia) of a basidiomycete related to Ustilago hordei (Hurek et al., 1997b); they were collected in rice field soil at the site in the Punjab of Pakistan, where formerly Kallar grass had been grown. The finding that culturable bacteria appear to persist in fungal resting stages but not in soil suggests that plantassociated fungi may play a role as shuttle vectors for reinfection of plants by these diazotrophic endophytes. However, Azonexus fungiphilus was thought to be exclusively fungusassociated (Reinhold-Hurek and Hurek, 2000) but was recently cultured from surface-sterilized roots of wild rice, as well (B. Reinhold-Hurek and Z. Y. Tan, unpublished observations). A new group of Azospira oryzae (Tan and Reinhold-Hurek, 2003), which was misnamed “Dechlorosoma suillum” has an entirely different habitat. It was isolated from a primary treatment lagoon of swine waste in the United States (Achenbach et al., 2001).

Isolation Since all known strains are diazotrophic, enrichment and isolation are done on nitrogen-free media. The procedures are the same as described for plant-associated Azoarcus spp.

Identification Azovibrio, Azonexus and Azospira are quite similar in physiological features but differ especially in cell morphology. Characters differentiating them from Azoarcus and other related genera are given in Table 1. More detailed phenotypic features are listed in Table 3. All three genera are diazotrophic. They do not use carbohydrates as sole carbon sources, but they do use a few organic and amino acids. Additional features are given below.

CHAPTER 3.2.18 Fig. 3. Microscopic images of cells of the genera Azovibrio, Azospira and Azonexus. Transmission electron (A) and phase contrast (B-E) images of Azovibrio restrictus S5b2T (A, B), Azospira oryzae 6a3T (C), and Azonexus fungiphilus BS5-8T (D, E). Cells were grown on N2 in semisolid medium and photos taken during exponential (B), stationary (E) and late stationary (C, D) growth phase. Bars, 1 µm (A), 5 µm (B-D) or 10 µm (E). From Reinhold-Hurek and Hurek (2000).

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

A

889

B

C

D

Azovibrio cells are motile, slightly curved rods of 0.6–0.8 × 1.5–3.6 µm (Fig. 3), and elongated cells occur very rarely in stationary phase cultures. Cells grow well at 37°C. The major cellular fatty acids are cis-9-16:1, 16:0, 18:1, 14:0 and 3-OH-10:1. They have no dissimilatory nitrate reductase. The only validly described species is A. restrictus; however, strain BS20-3 (isolated from black sclerotia) might represent another species. In contrast to A. restrictus, BS20-3 requires cobalamin; it is also unusual because it is strictly microaerobic with negligible colony growth (Reinhold-Hurek and Hurek, 2000). The type strain is A. restrictus S5b2 (= LMG9099) which has a G+C content of 64.8 mol% and was isolated from surface-sterilized roots of Kallar grass. Accession number of the 16S rDNA sequence is AF011349 (www.ncbi.nlm.nih.gov/ entrez/query.fcgi?db=nucleotide=search= AF011349). Azospira cells are thin curved rods of 0.4–0.6 × 1.1–2.5 µm (Fig. 3), and elongated cells with one to several spiral windings of up to 8 µm occur rarely in late-stationary phase nitrogen-fixing cultures. Cells are highly motile with corkscrewlike motion. Growth is optimum at 37°C or 40°C. The major cellular fatty acids are cis-9-16:1, 16:0, 18:1 and 3-OH-10:1. They have no dissimilatory nitrate reductase. Plant-associated strains can use oxygen and nitrate as terminal electron acceptors, while strains originating from wastewater (formerly [Dechlorosoma suillum]) can also use perchlorate. The only validly described species is Azospira oryzae. The type strain is

E

strain 6a3 (= LMG9096) which has a G+C content of 65.2 mol% and was isolated from surfacesterilized roots of Kallar grass. Accession number of the 16S rDNA sequence is AF011347 (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db= nucleotide=search=AF011347). Azonexus cells are highly motile, slightly curved rods of 0.6–0.8 × 1.5–4.0 µm; elongated straight to coiled cells of up to 50 µm length occur in the stationary phase of nitrogen-fixing cultures (Fig. 3). Cells grow well at 37°C and require cobalamin for growth. The only validly described species is Azonexus fungiphilus. The type strain is strain BS5-8 (= LMG 19178), which was isolated from resting stages (black sclerotia) of a basidiomycete related to Ustilago found in a rice field in Pakistan. Accession number of the 16S rDNA sequence is AF011350 (www.ncbi. nlm.nih.gov/entrez/query.fcgi?db=nucleotide= search=AF011350).

Preservation Preservation and storage are carried out as for plant-associated species of Azoarcus.

Physiology All three genera (even the perchlorate-reducing isolates from swine wastewater) are diazotrophic, showing microaerophilic nitrogen fixation (Tan and Reinhold-Hurek, 2003). As the plant-associated Azoarcus strains, cells require microaerobic conditions for nitrogen fixation

890

B. Rareinhold-Hurek and T. Hurek

and form subsurface pellicles due to microaerophilic growth in semisolid media. The optimum growth temperature is 37–40°C. Optimum pH is neutral, and the bacteria are not halophilic. All genera have a strictly respiratory type of metabolism, and no fermentative abilities have been observed. All genera use oxygen or nitrate as the terminal electron acceptor, and some strains of Azospira oryzae also use perchlorate. All are oxidase and catalase positive. Similarly to plantassociated Azoarcus species, organic acids, amino acids, and ethanol but no carbohydrates are utilized.

Genetics Extrachromosomal genetic elements have not been intensively studied in these genera. Transposon mutagenesis has been established for Azonexus fungiphilus (Gemmer and B. Reinhold-Hurek, unpublished), while Azospira oryza appears to be difficult to transform.

Literature Cited Achenbach, L. A., U. Michaelidou, R. A. Bruce, J. Fryman, and J. D. Coates. 2001. Dechloromonas agitata gen. nov., sp. nov. and Dechlorosoma suillum gen. nov., sp. nov., two novel environmentally dominant (per)chloratereducing bacteria and their phylogenetic position. Int. J. Syst. Evol. Microbiol. 51:527–533. Anders, H. J., A. Kaetzke, P. Kämpfer, W. Ludwig, and G. Fuchs. 1995. Taxonomic position of aromatic-degrading denitrifying pseudomonad strains K 172 and KB 740 and their description as new members of the genera Thauera, as Thauera aromatica sp. nov., and Azoarcus, as Azoarcus evansii sp. nov., respectively, members of the beta subclass of the Proteobacteria. Int. J. Syst. Bacteriol. 45:327–333. Beller, H. R., and A. M. Spormann. 1997. Anaerobic activation of toluene and o-xylene by addition to fumarate in denitrifying strain T. J. Bacteriol. 179:670–676. Boll, M., G. Fuchs, and J. Heider. 2002. Anaerobic oxidation of aromatic compounds and hydrocarbons. Curr. Opin. Chem. Biol. 6:604–611. Braun, K., and D. T. Gibson. 1984. Anaerobic degradation of 2-aminobenzoate (anthranilic acid) by denitrifying bacteria. Appl. Environ. Microbiol. 48:102–107. Chee-Sanford, J. C., W. F. Frost, M. R. Fries, J. Zhou, and J. M. Tiedje. 1996. Evidence for acetyl coenzyme A and cinnamoyl coenzyme A in the anaerobic toluene mineralization pathway in Azoarcus tolulyticus Tol-4. Appl. Environ. Microbiol. 62:964–973. Dörr, J., T. Hurek, and B. Reinhold-Hurek. 1998. Type IV pili are involved in plant-microbe and fungus-microbe interactions. Molec. Microbiol. 30:7–17. Egener, T., T. Hurek, and B. Reinhold-Hurek. 1999. Endophytic expression of nif genes of Azoarcus sp. strain BH72 in rice roots. Molec. Plant-Microb. Interact. 12:813–819. Egener, T., D. E. Martin, A. Sarkar, and B. Reinhold-Hurek. 2001. Role of a ferrodoxine gene cotranscribed with the nifHDK operon in N2 fixation and nitrogenase “switch

CHAPTER 3.2.18 off” of Azoarcus sp. strain BH72. J. Bacteriol. 183:3752– 3760. Engelhard, M., T. Hurek, and B. Reinhold-Hurek. 2000. Preferential occurrence of diazotrophic endophytes, Azoarcus spp., in wild rice species and land races of Oryza sativa in comparison with modern races. Environ. Microbiol. 2:131–141. Fries, M., J. Zhou, J. Chee-Sanford, and J. M. Tiedje. 1994. Isolation, characterization, and distribution of denitrifying toluene degraders from a variety of habitats. Appl. Environ. Microbiol. 60:2802–2810. Fries, M. R., J. J. Forney, and J. M. Tiedje. 1997. Phenol- and toluene-degrading microbial populations from an aquifer in which successful trichloroethene cometabolism occurred. Appl. Environ. Microbiol. 63:1523–1530. Harwood, C., G. Burchhardt, H. Herrmann, and G. Fuchs. 1998. Anaerobic metabolism of aromatic compounds via the benzoyl-CoA pathway. FEMS Microbiol. Rev. 22:439–458. Hurek, T., B. Reinhold, I. Fendrik, and E. G. Niemann. 1987. Root-zone-specific oxygen tolerance of Azospirillum spp. and diazotrophic rods closely associated with Kallar grass. Appl. Environ. Microbiol. 53:163–169. Hurek, T., B. Reinhold-Hurek, G. L. Turner, and F. J. Bergersen. 1994a. Augmented rates of respiration and efficient nitrogen fixation at nanomolar concentrations of dissolved O2 in hyperinduced Azoarcus sp. strain BH72. J. Bacteriol. 176:4726–4733. Hurek, T., B. Reinhold-Hurek, M. van Montagu, and E. Kellenberger. 1994b. Root colonization and systemic spreading of Azoarcus sp. strain BH72 in grasses. J. Bacteriol. 176:1913–1923. Hurek, T., and B. Reinhold-Hurek. 1995a. Identification of grass-associated and toluene-degrading diazotrophs, Azoarcus spp., by analyses of partial 16S ribosomal DNA sequences. Appl. Environ. Microbiol. 61:2257– 2261. Hurek, T., M. Van Montagu, E. Kellenberger, and B. Reinhold-Hurek. 1995b. Induction of complex intracytoplasmic membranes related to nitrogen fixation in Azoarcus sp. BH72. Molec. Microbiol. 18:225–236. Hurek, T., T. Egener, and B. Reinhold-Hurek. 1997a. Divergence in nitrogenases of Azoarcus spp., Proteobacteria of the β-subclass. J. Bacteriol. 179:4172–4178. Hurek, T., B. Wagner, and B. Reinhold-Hurek. 1997b. Identification of N2-fixing plant- and fungus-associated Azoarcus species by PCR-based genomic fingerprints. Appl. Environ. Microbiol. 63:4331–4339. Hurek, T., and B. Reinhold-Hurek. 1998. Interactions of Azoarcus sp. with rhizosphere fungi. In: A. Varma and B. Hock (Eds.) Mycorrhiza. Springer-Verlag. Berlin, Germany. 595–614. Hurek, T., Z. Tan, N. Mathan, T. Egener, M. Engelhard, P. Gyaneshwar, J. K. Ladha, and B. Reinhold-Hurek. 2000. Novel nitrogen-fixing bacteria associated with the root interior of rice. In:J. K. Ladha and P. M. Reddy (Eds.) The Quest for Nitrogen Fixation in Rice. International Rice Research Institute. Los Banos, The Philippines. 47– 62. Hurek, T., L. Handley, B. Reinhold-Hurek, and Y. Piché. 2002. Azoarcus grass endophytes contribute fixed nitrogen to the plant in an unculturable state. Molec. PlantMicrob. Interact. 15:233–242. Hurek, T., and B. Reinhold-Hurek. 2003. Azoarcus sp. strain BH72 as a model for nitrogen-fixing grass endophytes. J. Biotechnol. 106:169–178.

CHAPTER 3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus

Juretschko, S., A. Loy, A. Lehner, and M. Wagner. 2002. The microbial community composition of a nitrifyingdenitrifying activated sludge from an industrial sewage treatment plant analyzed by the full-cycle rRNA approach. Syst. Appl. Microbiol. 25:84–99. Juteau, P., R. Larocque, D. Rho, and A. LeDuy. 1999. Analysis of the relative abundance of different types of bacteria capable of toluene degradation in a compost biofilter. Appl. Microbiol. Biotechnol. 52:863–868. Krieger, C. J., W. Roseboom, S. P. Albracht, and A. M. Spormann. 2001. A stable organic free radical in anaerobic benzylsuccinate synthase of Azoarcus sp. strain T. J. Biol. Chem. 276:12924–12927. Laguerre, G., B. Bossand, and R. Bardin. 1987. Free-living dinitrogen-fixing bacteria isolated from petroleum refinery oily sludge. Appl. Environ. Microbiol. 53:1674– 1678. Macy, J. M., S. Rech, G. Auling, M. Dorsch, E. Stackebrandt, and L. I. Sly. 1993. Thauera selenatis gen. nov. sp. nov., a member of the beta subclass of Proteobacteria with a novel type of anaerobic respiration. Int. J. Syst. Bacteriol. 43:135–142. Mechichi, T., E. Stackebrandt, N. Gad’on, and G. Fuchs. 2002. Phylogenetic and metabolic diversity of bacteria degrading aromatic compounds under denitrifying conditions, and description of Thauera phenylacetica sp. nov., Thauera aminoaromatica sp. nov., and Azoarcus buckelii sp. nov. Arch. Microbiol. 178:26–35. Mohamed, M. E.-S. 2000. Biochemical and molecular characterization of phenylacetate-coenzyme A ligase, an enzyme catalyzing the first step in aerobic metabolism of phenylacetic acid in Azoarcus evansii. J. Bacteriol. 182:286–294. Mohamed, M. E., A. Zaar, C. Ebenau-Jehle, and G. Fuchs. 2001. Reinvestigation of a new type of aerobic benzoate metabolism in the proteobacterium Azoarcus evansii. J. Bacteriol. 183:1899–1908. Rabus, R., and F. Widdel. 1995. Anaerobic degradation of ethylbenzene and other aromatic hydrocarbons by new denitrifying bacteria. Arch. Microbiol. 163:96–103. Reinhold, B., T. Hurek, E.-G. Niemann, and I. Fendrik. 1986. Close association of Azospirillum and diazotrophic rods with different root zones of Kallar grass. Appl. Environ. Microbiol. 52:520–526. Reinhold-Hurek, B., T. Hurek, M. Gillis, B. Hoste, M. Vancanneyt, K. Kersters, and J. De Ley. 1993. Azoarcus gen. nov., nitrogen-fixing proteobacteria associated with roots of Kallar grass (Leptochloa fusca (L.) Kunth) and

891

description of two species Azoarcus indigens sp. nov. and Azoarcus communis sp. nov. Int. J. Syst. Bacteriol. 43:574–584. Reinhold-Hurek, B., and T. Hurek. 1998a. Interactions of gramineous plants with Azoarcus spp. and other diazotrophs: Identification, localization and perspectives to study their function. Crit. Rev. Plant Sci. 17:29–54. Reinhold-Hurek, B., and T. Hurek. 1998b. Life in grasses: Diazotrophic endophytes. Trends Microbiol. 6:139–144. Reinhold-Hurek, B., and T. Hurek. 2000. Reassessment of the taxonomic structure of the diazotrophic genus Azoarcus sensu lato and description of three new genera and species, Azovibrio restrictus gen. nov., sp. nov., Azospira oryzae gen. nov., sp. nov, and Azonexus funguphilus gen. nov., sp. nov. Int. J. Syst. Evol. Microbiol. 50:649–659. Rhee, S. K., G. M. Lee, J. H. Yoon, Y. H. Park, H. S. Bae, and S. T. Lee. 1997. Anaerobic and aerobic degradation of pyridine by a newly isolated denitrifying bacterium. Appl. Environ. Microbiol. 63:2578–2585. Song, B., M. M. Haggblom, J. Zhou, J. M. Tiedje, and N. J. Palleroni. 1999. Taxonomic characterization of denitrifying bacteria that degrade aromatic compounds and description of Azoarcus toluvorans sp. nov. and Azoarcus toluclasticus sp. nov. Int. J. Syst. Bacteriol. 49:1129– 1140. Springer, N., W. Ludwig, B. Philipp, and B. Schink. 1998. Azoarcus anaerobius sp. nov., a resorcinol-degrading, strictly anaerobic, denitifying bacterium. Int. J. Syst. Bacteriol. 48:953–956. Tan, Z. Y., and B. Reinhold-Hurek. 2003. Dechlorosoma suillum Achenbach et al. 2001 is a later subjective synonym of Azospira oryzae Reinhold-Hurek and Hurek 2000. Int. J. Syst. Evol. Microbiol. 53:1139–1142. Ueda, T., Y. Suga, N. Yahiro, and T. Matsuguchi. 1995. Remarkable N2-fixing bacterial diversity detected in rice roots by molecular evolutionary analysis of nifH gene sequences. J. Bacteriol. 177:1414–1417. Widdel, F., and F. Bak. 1992. Gram-negative mesophilic sulfate-reducing bacteria. In: A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes, 2nd ed. Springer. Berlin, Germany. 4:3352– 3378. Zhou, J., M. R. Fries, J. C. Chee-Sanford, and J. M. Tiedje. 1995. Phylogenetic analyses of a new group of denitrifiers capable of anaerobic growth on toluene and description of Azoarcus tolulyticus sp. nov. Int. J. Syst. Bacteriol. 45:500–506.

Index

A Absorption spectra Chromobacterium, 737 Rhodoferax, 596 Acetan production, Gluconacetobacter, 183–84 Acetate production Acetobacter, 176–77 Acetobacteraceae, 163 Acidomonas, 178 Asaia, 179–80 Gluconacetobacter, 180, 183 Kozakia, 190 phototrophic alphaproteobacteria, 55 Acetate utilization, phototrophic alphaproteobacteria, 56 Acetic acid bacteria. See Acetobacteraceae Acetic acid oxidation Acetobacter, 175–76 Acidomonas, 179 Gluconacetobacter, 181 Kozakia, 190 Acetobacter, 13, 151, 163–66, 168–70, 175–78. See also Gluconacetobacter applications of, 178 characteristics of, 168, 176–77 classification of, 167, 175–76 genetics of, 177–78 habitats of, 176 identification of, 176 isolation and cultivation of, 176 pathogenicity of, 178 taxonomy of, 175–76 Acetobacter aceti, 166, 168, 170–71, 174–78 Acetobacter acidophilus, 171 Acetobacter europaeus, 172 Acetobacter indonesiensis, 169 Acetobacter lovaniensis, 169, 173, 177 Acetobacter malorum, 177 Acetobacter oboediens, 172 Acetobacter orleanensis, 173 Acetobacter pasteurianus, 166, 168–73, 175–78, 731 Acetobacter peroxydans, 166, 176, 178 Acetobacter pomorum, 176–77 Acetobacter suboxydans, 365

Acetobacter tropicalis, 169 Acetobacter xylinum, 168, 171 Acetobacteraceae, 13, 163–90 Acetobacter, 13, 163–66, 175–78 Acidiphilium, 13 Acidomonas, 13, 163–66, 178–79 Asaia, 13, 163–66, 179–80 characteristics of, 168 classification of, 167 Gluconacetobacter, 13, 163–66, 180–85 Gluconobacter, 13, 163–66, 185–89 habitats and uses of, 163, 166–75 isolation, cultivation and preservation of, 175 Kozakia, 163–66, 189–90 morphology of, 163 phylogenetic tree of, 163–65 phylogeny of, 163–64 Rhodopila, 13 taxonomy of, 164–66 Acetone degradation, Xanthobacter, 308–9 Acetylmethylcarbinol production, Gluconobacter, 189 Achromobacter, 648, 678–89 applications of, 687–89 arsenite resistance by, 686 compound degradation by, 683–84 detection of, 686–87 heavy metal resistance by, 686 as human pathogens, 678–80 physiology of, 679 Achromobacter cycloclastes, 680–81, 687 Achromobacter denitrificans, 678 Achromobacter EST4002, 685 Achromobacter insolitus, 678 Achromobacter lyticus M497-1, 688 Achromobacter piechaudii, 678 Achromobacter ruhlandii, 678 Achromobacter sp. WM111, 686 Achromobacter spanius, 678 Achromobacter strain ABIV, 685 Achromobacter strain M-30-Y, 686 Achromobacter strain RS9, 685 Achromobacter xylosoxidans, 675, 678, 682–83, 686–87, 747 Achromobacter xylosoxidans subsp. denitrificans, 685 Achromobacter xylosoxidans subsp. xylosoxidans B3, 686 Achromopeptidase, 688

Acid pollution, Thiobacillus, 815 Acid production, Thiobacillus, 816, 819 Acidianus, 815 Acidiphilium, 13, 41, 163, 562, 565–66, 576, 580 Acidiphilium acidophilum, 819, 821 Acidiphilium rubrum, 571, 580 Acidisphaera rubrifaciens, 163 Acidithiobacillus, 18, 822–24 Acidithiobacillus albertis, 816, 822–23 Acidithiobacillus caldus, 823 Acidithiobacillus ferrooxidans, 815–16, 818–19, 822–24 Acidithiobacillus thiooxidans, 816, 822–24 Acidocella, 163 Acidomonas, 13, 163–66, 178–79 applications of, 179 characteristics of, 168, 179 classification of, 167 genetics of, 179 habitats of, 178 identification of, 178–79 isolation and cultivation of, 178 taxonomy of, 178 Acidomonas methanolica, 178–79 Acidophile Acidithiobacillus, 822–24 Beijerinckia, 151–52 Thiobacillus, 812 Acidovorax, 16, 723 Acidovorax facilis, 16 Acinetobacter, 20 Acremonium alternatum, 885 Actinobacillus, 19 Actinobacillus actinomycetemcomitans, 844–45 Actinomyces, 845 Activated sludge Achromobacter/Alcaligenes strains, 684–85 Alcaligenes, 677 Alcaligenes strains, 682 Azoarcus, 876 bulking of, 772 Comamonas, 726, 732 Leptothrix cholodnii, 768 Leptothrix lopholea, 768 Leptothrix-Sphaerotilus group, 763

894

Index

nitrite-oxidizing bacteria, 863, 865, 868 phototrophic betaproteobacteria, 596 Sphaerotilus, 772–73 Adenylate cyclase toxin, 662–64, 666, 668 Bordetella, 658, 660 Adherence Bordetella, 656–57, 667 Eikenella corrodens, 840 Adhesins Bordetella, 655–56, 665 Opa proteins as, 611 as toxins, 657 Aerobic phototrophic bacteria, 562–80 applications of, 580 ecology of, 580 genetics of, 578–80 habitats of, 565–66 identification of, 568–71 isolation of, 566–68 physiology of, 572–78 oxygen dependence and heterotrophic metabolism, 575–78 photosynthetic potential, 572–75 preservation of, 571 properties of, 563–64, 567 taxonomy, phylogeny and origin, 562–65 Aerocavin, 741 Aerocyanidin, 741 Aeromonadaceae, 19–20 Aeromonas, 19 Aeromonas, 19, 740 Aeromonas salmonicida, 19 Afipia, 15 African tick-bite fever, Rickettsia, 503 Agave habitat, Zymomonas, 201 Agracin sensitivity, Agrobacterium, 107, 109 Agrobacterium, 13–15, 91–110, 151, 315, 321, 324, 350, 365, 467–68, 659, 747, 749 applications of, 109–10 characteristics of, 94 disease from, 91–110, 105–9 binding to plant, 107–9 control of, 106–9 ecology of, 105 genetics of, 96–105 chromosomal, 97–101 general, 96–97 habitats of, 92 identification of, 92–94 isolation of, 92 morphology of, 94 phylogeny of, 91 physiology of, 94–96 preservation of, 94 taxonomy of, 91–92 Agrobacterium radiobacter, 91, 107, 109, 468 Agrobacterium rhizogenes, 91–92, 95, 101, 106, 109–10

Agrobacterium rubi, 91 Agrobacterium tumefaciens, 28, 91–92, 94–103, 105–10, 127–28, 130, 324, 371–72 Agrobacterium vitis, 91–92, 101, 104–6 Albamycin sensitivity, Methylobacterium, 262 Alcaligenaceae, 16–17, 675, 678, 689, 747 Alcaligenes, 16, 648, 675–77, 723, 754, 849 applications of, 687–89 arsenite resistance by, 686 compound degradation by, 682–86 denitrification by, 680–82 detection of, 686–87 heavy metal resistance by, 686 as human pathogens, 678–80 physiology of, 679 strains of, 681–88 Alcaligenes defragrans, 675, 677, 684 Alcaligenes denitrificans, 681, 683, 685–87 Alcaligenes denitrificans subsp. denitrificans, 689 Alcaligenes denitrificans subsp. xylosoxidans, 678, 682 Alcaligenes faecalis, 365, 675, 677, 680–83, 686–87, 689, 751, 868 Alcaligenes faecalis subsp. faecalis, 675, 677 Alcaligenes faecalis subsp. parafaecalis, 675, 677, 688 Alcaligenes latus, 675, 758–59 Alcaligenes xylosoxidans, 680–87, 850 Alcaligenes xylosoxidans subsp. denitrificans, 685, 688 Alcaligenes xylosoxidans subsp. xylosoxidans, 688 Alcalilimnicola, 19 Alcohol dehydrogenase isoenzymes, Zymomonas, 206–7 Alcohol utilization Acidomonas, 179 Agrobacterium, 91–92 Azospirillum, 115 Gluconobacter, 187 Herbaspirillum, 145–46 Kozakia, 190 Methylobacterium, 257 Paracoccus, 238, 240–43 phototrophic alphaproteobacteria, 56 Xanthobacter, 305–6 Algal habitat Aquaspirillum, 711 dimorphic prosthecate bacteria, 76–77, 86 Alkane degradation, Xanthobacter, 307–8 Alkanesulfonate degradation, Achromobacter/Alcaligenes strains, 684 Alkene degradation, Xanthobacter, 307–8 Allorhizobium, 14–15 Alphaproteobacteria, 3–29

Acetobacteraceae, 13 Anaplasmataceae, 13 Bartonellaceae, 14–15 Brucellaceae, 14–15 Caulobacteraceae, 14 ecology of, 9–15 Ehrlichiaceae, 13 group 1, 43–44 group 2, 44 group 3, 44 Hyphomicrobiaceae, 15, 290 morphology of, 9–15 orders and families of, 12 phototrophic, 41–59 phylogeny of, 9–15 physiology of, 9–15 Rhizobiaceae, 14–15 Rhizobiales, 294 Rhodospirillaceae, 13–14 Rickettsiaceae, 13 Rickettsiales, 457–64 Sphingomonadaceae, 14 Alteromonadaceae, 19–20 Alteromonas, 737 Alteromonas luteoviolacea, 740, 742 Alysiella, 18, 828–38 ecology of, 837–38 habitat of, 832–34 identification of, 835–37 isolation of, 834–35 morphology of, 828, 830, 834–36 phylogeny of, 828–32 preservation of, 837 taxonomy of, 832 traits of, 832 Alysiella filiformis, 828, 830–33, 836–37 Amaricoccus, 14 Amino acid synthesis Achromobacter/Alcaligenes strains, 687 Comamonas, 729 Thiobacillus, 819 Amino acid utilization aerobic phototrophic bacteria, 576 Brucella, 362, 366 dimorphic prosthecate bacteria, 82–83 epsilonproteobacteria, 24 Neisseria, 608 Paracoccus, 240–43 phototrophic alphaproteobacteria, 56 D-Aminoacylases, Achromobacter/Alcaligenes strains, 687 1-Aminocyclopropane-1-carboxylate deaminase, Alcaligenes strains, 682 Aminoglycoside sensitivity Brucella, 428–29 dimorphic prosthecate bacteria, 74 Ammonia assimilation, phototrophic betaproteobacteria, 598 Ammonia-oxidizing bacteria, 778–806, 862, 864, 866 characteristics of, 779–86

Index detection of, 794–800 classical techniques, 794–95 immunological techniques, 795 molecular techniques, 796–800 distribution in nature, 800–803 betaproteobacteria, 801–3 gammaproteobacteria, 803 enrichment, isolation, and maintenance of, 778–79 environmental populations, 803–6 genera of, 779–80 habitats of, 781 morphology of, 780, 783–86 phylogenetic tree of, 787–90 phylogeny of, 786–94 species of, 780–86 Ammonium utilization Azospirillum, 129 dimorphic prosthecate bacteria, 74 Ampicillin resistance, Xanthobacter, 303 Amylosucrase, Neisseria, 624 Anaerobiospirillum, 19–20 Anaplasma, 13, 320, 323, 460–61, 473, 493–94, 507–12, 547 ecology of, 510 epidemiology of, 510–11 genetics of, 509–10 identification of, 508–9 isolation of, 508 phylogeny of, 507–8 physiology of, 509 preservation of, 509 Anaplasma bovis, 493, 508 Anaplasma centrale, 511 Anaplasma marginale, 460–61, 493, 508–12 Anaplasma ovis, 508 Anaplasma phagocytophila, 493–94, 508–10 Anaplasma platys, 508–9, 511, 514 Anaplasmataceae, 13, 493 Anaplasma, 13, 493 Ehrlichia, 493 habitats of, 508 morphology of, 493 Neorickettsia, 493 phylogenetic tree of, 508 taxonomy of, 508 Wolbachia, 493, 547 Anaplasmosis Anaplasma, 508–12 control and prevention of, 511 vaccine for, 511–12 Ancalomicrobium, 65–70 characteristics of, 69 cultivation and maintenance of, 68–69 habitats of, 65–66 identification of, 69–70 isolation of, 66–68 morphology of, 65 phylogenetic tree of, 67 species of, 67 taxonomy of, 65 Ancalomicrobium adetum, 65–66, 68–69 Angiococcus, 22

Angiogenic factor, Bartonella, 472–73 AniA, 613 Neisseria, 607 Aniline degradation, Comamonas, 729 Animal disease Aeromonadaceae, 19 Anaplasma, 511 Anaplasmataceae, 493 Bartonella, 462, 467 Betaproteobacteria, 16 Brucella, 315, 335–38, 372–429 Campylobacteraceae, 24–25 Cardiobacteriaceae, 20 Enterobacteriaceae, 19 Francisella, 20 Gammaproteobacteria, 18 Neisseriaceae, 18 Pasteurellaceae, 19 Piscirickettsia, 20 Animal habitats Neisseria, 603 Simonsiella, 829 Antarctobacter, 14 Antibiotic production Chromobacterium, 741 Myxobacteria, 22 Antibiotic resistance/sensitivity Achromobacter/Alcaligenes strains, 680, 689 Azoarcus, 886 Bartonella, 484 Bordetella, 653 Brucella, 321, 350, 411–12, 428–29 Burkholderia, 852, 856 Chromobacterium, 739 dimorphic prosthecate bacteria, 74 Methylobacterium, 262 Neisseria, 624–25, 626, 630, 632 Rickettsia, 506 Spirillum, 707 Xanthobacter, 303 Antibiotic treatment gonorrhea, 632 meningococcal disease, 632 pertussis, 651, 653 Antibody response, to pertussis, 665–66 Antigenic structure, Eikenella corrodens, 842 Antigens Burkholderia, 854, 855 Neisseria, 617 Antimicrobial peptides, 664 Bordetella, 661–62 Antimicrobial susceptibility, Eikenella corrodens, 844 Antipyrin degradation, Phenylobacterium, 250–56 Appendicitis, Bilophila, 23 Appendix, Comamonas, 726 Apple habitat, Acetobacteraceae, 172–73 Applications Acetobacter, 178 Achromobacter/Alcaligenes strains, 687–89

895

Acidomonas, 179 aerobic phototrophic bacteria, 580 Agrobacterium, 109–10 Azospirillum, 132–34 Beijerinckia, 161 Brucella, 429–31 Chromobacterium, 740–41 Comamonas, 732 Coxiella, 541 Derxia, 756 dimorphic prosthecate bacteria, 85–86 Gluconacetobacter, 185 Gluconobacter, 188–89 Herbaspirillum, 147 Janthinobacterium, 743 manganese-oxidizing bacteria, 229 methanotrophs, 284 Methylobacterium, 263 Ochrobactrum, 749 Paracoccus, 246 phototrophic alphaproteobacteria, 58–59 phototrophic betaproteobacteria, 599 Phyllobacterium, 749 Proteobacteria, 12 Spirillum, 707–8 Wolbachia, 516, 555–56 Xanthobacter, 307–9 Aquabacter, 290, 294 Aquabacter spiritensis, 294 Aquabacterium, 758 Aquaspirillum, 701, 703, 710–21, 723 differentiating characteristics of, 712–13 enrichment of, 711–18 habitat of, 711 identification of, 719–21 isolation of, 711–19 maintenance of, 711–18 morphology of, 719 motility of, 719 preservation of, 718–19 rRNA superfamily III species of, 719–20 rRNA superfamily IV species of, 720–21 taxonomy of, 719–21 type strains of, 714 Aquaspirillum anulus, 720 Aquaspirillum aquaticum, 711, 717, 719, 720, 725–26 Aquaspirillum autotrophicum, 710, 711, 717, 720, 742–43 Aquaspirillum bengal, 710, 717, 719 Aquaspirillum delicatum, 710, 719, 720 Aquaspirillum dispar, 719, 720 Aquaspirillum fasciculus, 710, 716, 719, 720 Aquaspirillum giesbergeri, 720 Aquaspirillum gracile, 710, 716, 719, 720 Aquaspirillum itersonii, 141, 703, 710, 719, 720 Aquaspirillum itersonii subsp. itersonii, 720

896

Index

Aquaspirillum itersonii subsp. nipponicum, 718, 720 Aquaspirillum itersonii subsp. vulgatum, 720 Aquaspirillum magnetotacticum, 710, 717, 718, 719, 720 Aquaspirillum metamorphum, 715, 720 Aquaspirillum peregrinum, 719, 720 Aquaspirillum peregrinum subsp. integrum, 718 Aquaspirillum peregrinum subsp. peregrinum, 710 Aquaspirillum polymorphum, 710, 719, 720 Aquaspirillum psychrophilum, 710, 718, 720 Aquaspirillum putridiconchylium, 720 Aquaspirillum serpens, 710, 716, 717, 719, 720 Aquaspirillum sinuosum, 720 Aquaspirillum voronezhense, 701 Archangiaceae, 22 Archangium, 22 Arcobacter, 24–25 Arcobacter nitrofigilis, 25 Aromatic amine deamination, Alcaligenes strains, 682 Aromatic amine dehydrogenase, Alcaligenes strains, 682 Aromatic compound degradation Achromobacter/Alcaligenes strains, 683–84 Azoarcus, 873, 886 Comamonas, 728 phototrophic betaproteobacteria, 598 Arphamenines A and B, 741 Arsenite resistance, Achromobacter/Alcaligenes strains, 686 Arthrobacter globiformis, 731 Arthropod habitat Alcaligenes, 677 Anaplasma, 508–9 Ancalomicrobium, 66 Bartonella, 479 Coxiella, 530, 536 dimorphic prosthecate bacteria, 77 Ehrlichia, 512–13 Prosthecomicrobium, 66 Rickettsia, 496, 500 Wolbachia, 515–16, 547, 550–51, 553 Asaia, 13, 163–66, 172–73, 179–80 characteristics of, 168, 180 classification of, 167 habitats of, 180 identification of, 180 isolation and cultivation of, 180 taxonomy of, 180 Asaia bogorensis, 180, 190 Asaia krungthepensis, 180 Asaia siamensis, 180 Asticcacaulis, 72–86 Asticcacaulis biprosthecum, 74, 81, 84

Asticcacaulis excentricus, 74, 81, 84 ATP synthesis, Brucella, 367 Atrophic rhinitis, 651, 664 Autophagosomes, Brucella, 407–8 Autotransporter proteins, Rickettsia, 498–99 Auxotyping, Neisseria, 628 Azo dye degradation, Pigmentiphaga kullae, 689 Azoarcus, 17, 126, 873–88 characteristics of, 878 ecology of, 886–87 genetics of, 886 habitat of, 875 identification of, 877–85 isolation of, 876–77 phylogeny of, 873–75 physiology of, 885–86 preservation of, 885 species of, 880–83 taxonomy of, 875 Azoarcus anaerobius, 873, 875, 877, 879, 885–86 Azoarcus buckelii, 873, 875, 877, 879, 885 Azoarcus communis, 873, 875, 877, 879–86 Azoarcus evansii, 873, 875–77, 884–86 Azoarcus indigens, 873, 875, 877, 879, 885–87 Azoarcus sp. strain BH72, 873, 875, 877, 879, 885–87 Azoarcus toluclasticus, 873, 875, 877, 884–85 Azoarcus tolulyticus, 873, 875, 877, 884–86 Azoarcus toluvorans, 873, 875, 877, 884–85 Azoarcus/Thauera branch, 873–75, 888 Azomonas, 21, 154–55, 751, 753–54 Azomonas agilis, 160 Azonexus, 873, 877, 887–90 characteristics of, 878 genetics of, 890 identification of, 888–89 isolation of, 888 morphology of, 889 phylogeny of, 888 physiology of, 889–90 preservation of, 889 species of, 881–83 taxonomy of, 888 Azonexus fungiphilus, 887–90 Azorhizobium, 290, 294 Azorhizobium caulinodans, 294 Azospira, 17, 873, 877, 887–90 characteristics of, 878 genetics of, 890 identification of, 888–89 isolation of, 888 morphology of, 889 phylogeny of, 888 physiology of, 889–90 preservation of, 889 species of, 881–83 taxonomy of, 888 Azospira oryzae, 887–90

Azospirillum, 42, 115–34, 141, 151, 753–54 applications of, 132–34 characteristics of, 124 cultivation of, 122–23 habitats and ecology of, 116–20 history of, 115 identification of, 123–26 morphology of, 115, 121 phylogenetic tree of, 116 physiology of, 126–32 ammonium uptake and assimilation, 129–30 nitrogen fixation, 129 osmotolerance, 131 plant growth regulating substances, 130–31 plant root colonization, 127–28 siderophores, 131–32 preservation of, 123 species of, 118, 124 taxonomy of, 115–16 Azospirillum amazonense, 115–16, 121, 123, 125–26, 128–29, 131–32 Azospirillum brasilense, 115–16, 118–20, 123, 125, 127–33, 146–47, 720 Azospirillum doebereinerae, 115–16, 120, 124–25 Azospirillum halopraeferens, 115–16, 121–22, 125, 131 Azospirillum irakense, 115–16, 122–23, 126, 131–33 Azospirillum largimobile, 116, 122, 125, 132 Azospirillum lipoferum, 115–16, 118, 120, 122–25, 128–29, 131–32, 754 Azotobacter, 21, 154–56, 751, 753–54, 756 Azotobacter acida. See Beijerinckia indica Azotobacter chroococcum, 160, 275 Azotobacter vinelandii, 146, 160 Azotomonas insolita, 751 Azovibrio, 873, 877, 887–90 characteristics of, 878 genetics of, 890 identification of, 888–89 isolation of, 888 morphology of, 889 phylogeny of, 888 physiology of, 889–90 preservation of, 889 species of, 881–83 taxonomy of, 888 Azovibrio restrictus, 888, 889 Azurins, 681

B Bacillary angiomatosis, Bartonella, 480–81, 483–84 Bacillary peliosis, Bartonella, 481, 483–84 Bacillus extorquens, 257 Bacillus faecalis alcaligenes, 723

Index Bacillus subtilis, 861 Bacitracin resistance, Methylobacterium, 262 Bacteremia Bartonella, 462, 482–84 Coxiella, 536–37 Bacteriochlorophyll aerobic phototrophic bacteria, 562, 571, 575 Alphaproteobacteria, 9, 13 Bacteriochlorophyll a aerobic phototrophic bacteria, 562, 566 Methylobacterium, 263 phototrophic alphaproteobacteria, 49, 54 Bacteriochlorophyll b, phototrophic alphaproteobacteria, 50, 54 Bacteriochlorophyll β, phototrophic alphaproteobacteria, 50 Bacteriophage Acidomonas, 179 Brucella, 346–48, 359 Neisseria, 626 Bacteriophage production, Bartonella, 471–73 Bacteriovorax, 23 Bacteroides corrodens, 840 Bacteroides ureolyticus, 840 Balneatrix alpica, 21 Bartonella, 15, 315, 320, 323, 462, 467–85, 493 disease from, 480–84 antimicrobial therapy, 484 clinical presentation, 480–83 diagnosis, 483–84 epidemiology of, 479–80 genetics of, 478–79 habitats of, 469 identification of, 469–71 immunity from, 484–85 isolation of, 469 morphology of, 467, 470 pathogenesis host adherence, 474–75 host colonization, 473–74 host invasion, 475–78 phylogenetic tree of, 468 phylogeny of, 467 physiology of, 471–78 angiogenic factor, 472–73 cell structure, 471 enzymes, 473 extracellular products, 471–72 growth and metabolism, 471 pathogenesis, 473–78 preservation of, 471 taxonomy of, 467–69 Bartonella alsatica, 462 Bartonella bacilliformis, 15, 462, 467, 469, 471–75, 477–79, 482, 484 Bartonella clarridgeiae, 462, 470–71, 474, 480, 484 Bartonella doshiae, 462 Bartonella elizabethae, 462, 470–72, 475, 478–80, 484 Bartonella grahamhii, 462 Bartonella henselae, 15, 462, 467–76, 478–81, 483–85

Bartonella koehlerae, 462 Bartonella peromysci, 462 Bartonella quintana, 15, 462, 467–73, 475, 477–80, 482–85 Bartonella talpae, 462 Bartonella taylorii, 462 Bartonella tribocorum, 462, 478 Bartonella vinsonii, 471, 478 Bartonella vinsonii subsp. arupensis, 462 Bartonella vinsonii subsp. berkhoffii, 462 Bartonella vinsonii subsp. vinsonii, 462 Bartonella washoensis, 462 Bartonella weissii, 462 Bartonellaceae, 14–15, 467–68 Bartonella, 15, 467–68 Bartonellae, 462 Bartonellosis, Bartonella, 479–80, 484 Basidiomycete Azoarcus, 887 Azonexus, 888 Bdellovibrio, 23 Bdellovibrio group, 23 Bacteriovorax, 23 Bdellovibrio, 23 Bee hive environment Acetobacteraceae, 173–74 Gluconobacter, 186 Simonsiella, 832 Zymomonas, 201 Beer Acetobacter, 176 Acetobacteraceae, 172 Gluconobacter, 186 Zymomonas, 201 Beggiatoa, 20, 815 Beijerinckia, 15, 151–61, 751–56 applications of, 161 cultivation of, 154 habitats of, 151–52 identification of, 154–57 isolation of, 152–54 life cycle of, 155 morphology of, 151–52, 154–59 physiology of, 159–60 preservation of, 154 species differentiation, 157–59 Beijerinckia derxii, 154, 157, 159 Beijerinckia derxii subsp. derxii, 159 Beijerinckia derxii subsp. venezuelae, 159 Beijerinckia fluminensis, 154, 156–59 Beijerinckia indica, 152–54, 156–59 Beijerinckia indica subsp. indica, 158 Beijerinckia indica subsp. lacticogenes, 156, 158 Beijerinckia indica var. alba, 158 Beijerinckia mobilis, 154–58 Benzoate degradation, Azoarcus, 886 Betaproteobacteria, 3–29. See also Phototrophic betaproteobacteria Alcaligenaceae, 16–17 ammonia-oxidizing bacteria, 783–86

897

Comamonadaceae, 16–17 ecology of, 15–18 morphology of, 15–18 Neisseriaceae, 18 orders and families of, 16 phototrophic, 593–99 phylogeny of, 15–18 physiology of, 15–18 sulfur oxidizing bacteria of, 812–14, 821–22 Bilophila, 23 Bilophila wadsworthia, 23 Biochemistry Acetobacter, 176–77 Asaia, 180 Bartonella, 470 Cardiobacterium hominis, 844–45 Derxia, 755–56 Eikenella corrodens, 842 Gluconobacter, 186–87 Kozakia, 190 Neisseria, 606 Xanthobacter, 301–3 Biocontrol, Burkholderia, 848 Biodegradation Azoarcus, 886 Betaproteobacteria, 15–17 nitrite-oxidizing bacteria, 868–69 Biofilms Leptothrix-Sphaerotilus group, 763 nitrite-oxidizing bacteria, 863–65, 867 Biology, for speciation, 334–37 Bioremediation aerobic phototrophic bacteria, 580 Burkholderia, 848 Comamonas, 732 methanotrophs, 266, 284 Methylobacterium, 263 Pseudomonas, 21 Xanthobacter, 308–9 Biosynthesis, Neisseria, 620–21 Biphenyl degradation Achromobacter/Alcaligenes strains, 683 Comamonas, 730 Biphenyl/chlorobiphenyl dioxygenase, Comamonas, 730 Bird habitat Bordetella, 651 Coxiella, 530 Blastochloris, 44 Blastochloris sulfoviridis, 44, 46, 50, 54, 57 Blastochloris viridis, 44, 50, 56 Blood Achromobacter, 678 Alcaligenes, 677 Aquaspirillum, 711 Comamonas, 726 Ochrobactrum, 748 Bordetella, 16, 648–68, 675, 678 autotransporter protein family in, 656–57 description of, 648 diseases from, 650–51, 651 ecophysiology of, 653–62

898

Index

evolution of, 650 habitats of, 651–53 immunity to, 664–66 intracellular state of, 667–68 isolation of, 651–53 LPS of, 661–62 phylogeny of, 649–50 significance of, 650–51 species of, 648–49 toxins of, 658–61 vaccine resistance of, 666–67 virulence factors of, 656–58, 667–68 Bordetella avium, 648, 651, 655, 660 Bordetella bronchiseptica, 648–50, 653–55, 661–62, 668 Bordetella bronchiseptica cluster, 648, 652, 660–61, 663–64 characterization of, 649–50 virulence factors of, 667–68 Bordetella hinzii, 648 Bordetella holmesii, 648–49 Bordetella parapertussis, 648–51, 661–62, 667 Bordetella pertussis, 16, 351, 648–56, 659–60, 662–64, 666–68 toxins of, 658–61 transcriptional responses to, 664 virulence factors of, 656–58 Bordetella petrii, 648 Bordetella trematum, 648 Boutonneuse fever, Rickettsia, 503 Bovine habitat Anaplasma, 510–11 Anaplasmataceae, 493 Bartonella, 469 Brucella, 335, 372–82, 394, 429 Campylobacteraceae, 24–25 Coxiella, 530, 536 Succinivibrionaceae, 19–20 Brachymonas, 723 Brackiella oedipodis, 16 Bradyrhizobium, 13, 15, 320, 864 Bradyrhizobium japonicum, 754 Brenneria, 19 Brevundimonas, 14, 18 Brucella, 9, 13, 315–431, 467–68, 747, 845 applications of, 429–31 as adjuvants, 431 antitumoral and antiviral activity, 430 as biological weapon, 431 foreign antigen delivery, 430 as immunological tools, 431 characteristics of, 339 classification of, 322–23, 344–46 disease from, 381–429 control of infection, 411–15 diagnosis of, 415–28 pathobiology of, 387–411 pathology of, 381–87 treatment of, 428–29 epidemiology of, 372–81 control, eradication and prevention, 380 geographical distribution of, 373–75 host infection, 375

vaccines and vaccination, 375–80 zoonosis, 380–81 genetics of, 327–28, 368–72 chromosomes, 368–71 genome, 368 habitats of, 315, 335, 337, 349 identification of, 325–26, 341–48 brucellaphages, 346–48 cellular and colonial characteristics, 341–44 identification and typing, 344–46 isolation of, 337–41 phylogenetic tree of, 316–17, 321 phylogeny of, 315–30 ancestor-descendant relationships, 315–20 gene acquisition, 330 genome evolution, 321–24 speciation of, 320 virulence structure derivation, 324–29 physiology of, 349–68 metabolism, 361–68 structure, 349–61 preservation of, 348–49 taxonomy of, 330–37 genetic composition of, 332–33 species definition, 333–37 Brucella abortus, 320, 324, 329–31, 335–39, 341, 343, 345, 348–56, 359–68, 370–71, 374, 377–80, 382–85, 388, 390–97, 400–405, 409, 412–13, 417–18, 422, 424–25, 429–30 Brucella canis, 320, 330–31, 334–37, 339, 344–45, 353–54, 359, 365–66, 368, 374, 380, 382, 385, 390, 415, 418, 426–27 Brucella melitensis, 28, 320, 330–31, 335–39, 341, 343–45, 348, 352–54, 356, 359–60, 363–68, 372–74, 376–77, 379–80, 382–83, 385, 388, 408, 413, 418, 422, 425–26, 429–31 Brucella neotomae, 330–31, 335–37, 339, 345, 348, 368, 374, 380, 382, 385 Brucella ovis, 42, 330–31, 334–39, 341, 343, 345, 353–54, 365–66, 368, 371, 374, 379–80, 382, 385, 390, 415, 418, 426–27 Brucella suis, 320–21, 330–31, 335–39, 344–45, 348, 364–65, 368, 370, 372, 374, 378, 380, 382, 385, 388, 418, 431 Brucellaceae, 14–15, 840 Brucellosis Brucella, 372–75, 381–429 control of infection, 411–15 antibody response, 411–12 T cell response, 412–15 diagnosis of, 415–28 in animals, 416–17 antibody detecting tests, 421–24 cell-mediated immunity tests, 425–28 in humans, 415–16

immunological tests, 417–21 pathobiology of, 387–411 host invasion, 387 replication in host cell, 393–401 survival outside host cells, 387–93 virulence mechanisms, 401–11 pathology of, 381–87 in animals, 382, 385–86 in fetus, 382–85 in humans, 385 male infections, 385 outcome and self cure, 386–87 treatment of, 428–29 in animals, 429 in humans, 428–29 Buchnera, 19, 463 Buchnera aphidicola, 19, 28, 463 Budding dimorphic prosthecate bacteria, 73, 82 Methylobacterium, 261 Methylosinus, 280, 282 phototrophic alphaproteobacteria, 42–44 Seliberia, 585 Burkholderia, 16–18, 21, 848–56 disease of, 852–53 epidemiology of, 851–52 habitat of, 850 identification of, 850–51 isolation of, 850 pathogenicity of, 853–56 taxonomy of, 849–50 treatment of, 856 Burkholderia caryophilli, 849 Burkholderia cepacia, 16–17, 28, 848–54, 856 Burkholderia cepacia complex, 678, 849–52 Burkholderia gladioli, 849 Burkholderia mallei, 16, 848–49, 851–53, 855–56 Burkholderia multivorans, 850–52 Burkholderia picketii, 849 Burkholderia pseudomallei, 16, 848–56 Burkholderia solanaceareum, 849 Burkholderia stabilis, 849–52 Burkholderia thailandensis, 848–49 Burkholderia vietnamensis, 849–52

C Cadmium resistance Achromobacter/Alcaligenes strains, 686 Comamonas, 731 Caedibacter, 462 Caedibacter caryophila, 462 Calvin cycle, Paracoccus, 235 Campylobacter, 24–25, 703 Campylobacter jejuni, 24, 703 Campylobacteraceae, 24–25 Arcobacter, 24–25 Campylobacter, 24–25 Dehalospirillum, 24–25 Sulfurospirillum, 25

Index Canine habitat Anaplasma, 511 Anaplasmataceae, 494 Bartonella, 469, 479 Ehrlichia, 460 Neorickettsia, 461 Succinivibrionaceae, 19–20 Canine monocytotropic ehrlichiosis, Ehrlichia, 514–15 Capnogtophaga, 844 Caprine habitat Brucella, 372–82, 384, 394 Coxiella, 530, 536 Carbofurane degradation, Achromobacter/Alcaligenes strains, 686 Carbohydrate metabolism, Neisseria, 608 Carbohydrate transport, Zymomonas, 203–4 Carbon dioxide production, phototrophic alphaproteobacteria, 55 Carbon dioxide requirement Brucella, 366–67 Neisseria, 607–8 Carbon dioxide utilization ammonia-oxidizing bacteria, 778 phototrophic alphaproteobacteria, 56 Xanthobacter, 304–5 Cardiobacteriaceae, 20 Cardiobacterium, 20 Dichelobacter, 20 Suttonella, 20 Cardiobacterium, 20 Cardiobacterium hominis, 20, 844–45 Carnimonas, 20 Carotenoids aerobic phototrophic bacteria, 570–71, 575 Azospirillum, 125 phototrophic alphaproteobacteria, 44, 49 Rhodocyclus, 594–96 Caryophanon, 835 Cat flea typhus, Rickettsia, 503–4 Cat scratch disease, Bartonella, 462, 480, 483–84 Catalase production Brucella, 368 Coxiella, 541 dimorphic prosthecate bacteria, 74 Methylocystis, 280 Methylosinus, 280 Neisseria, 607 Catechol degradation, Comamonas, 728, 730 Catenococcus thiocyclus, 813, 817 Catheters, Comamonas, 726 Cats. See Feline habitat Caulobacter, 65, 72–86, 315 Caulobacter crescentus, 84, 370 Caulobacter leidyi, 77, 84 Caulobacter vibrioides, 83–84 Caulobacteraceae, 14 Brevundimonas, 14 CDC group Vd, 747, 748

Cell wall composition Bartonella, 471 Coxiella, 531 dimorphic prosthecate bacteria, 82 Methylococcus, 275 Orientia, 506 Rickettsia, 457 Xanthobacter, 302 Zymomonas, 203 Cell wall degradation, Achromobacter/Alcaligenes strains, 688 Cellulose production Agrobacterium, 97, 109 Gluconacetobacter, 183–85 Cellvibrio, 21 Cepabactin, Burkholderia, 853 Cepacia syndrome, 848 Cephalothin resistance, Methylobacterium, 262 Chemoprophylaxis, for meningococcal disease, 632 Chemostat culture, Thiobacillus, 817–19 Chemotaxis Agrobacterium, 96, 106 Azospirillum, 127 Xanthobacter, 301 Chicken habitat Alysiella, 832 Simonsiella, 832 Chlamydia, 463 Chlamydia pecorum, 463 Chlamydia pneumoniae, 463 Chlamydia psittaci, 463 Chlamydia trachomatis, 463 Chlamydiae, 463 Chlamydia, 463 morphology of, 463 Chloramphenicol resistance/ sensitivity Rickettsia, 506 Xanthobacter, 303 Chlorhexidine solutions, Achromobacter, 678 Chloridazon degradation, Phenylobacterium, 250–56 Chlorinated compound degradation, Achromobacter/Alcaligenes strains, 685 Chromate-reducing bacteria, Comamonas, 731 Chromatiaceae, 18–19 Chromatibacteria, 15 Chromatium, 703 Chromobacter violaceum, 851 Chromobacteriosis, 739 Chromobacterium, 18, 720, 737–41, 747 applications of, 740–41 differentiating characteristics of, 738 enrichment of, 739–40 habitat of, 738 identification of, 740 isolation of, 739–40 morphology of, 738

899

motility of, 738 preservation of, 740 Chromobacterium amethystinum, 741 Chromobacterium fluviatile, 737, 739–40 Chromobacterium lividum, 737 Chromobacterium membranaceum, 741 Chromobacterium violaceum, 737–42, 751 Chromohalobacter, 20 Chromosomes Agrobacterium, 91, 96–101, 110 Brucella, 321–24, 368–71 Coxiella, 535 Paracoccus, 245–46 Chronic granulomatous disease, 852 Cider Acetobacter, 176 Acetobacteraceae, 169–70 Gluconobacter, 186 Zymomonas, 201 Citrate utilization, phototrophic alphaproteobacteria, 56 Citric acid cycle, Phenylobacterium, 255 Citromicrobium, 562, 565, 571, 576 Citromicrobium bathyomarinum, 565–66, 572–73 Clarithromycin sensitivity, Orientia, 507 Clostridium, 688 Clostridium beijerinckii, 688 Cocoa wine Acetobacter, 176 Acetobacteraceae, 169 Gluconobacter, 186 Zymomonas, 201 Co-divergence hypothesis, 830 Coenzyme M, Xanthobacter, 308 Coffee plant habitat Acetobacteraceae, 173 Gluconacetobacter, 181, 184 Collectins, 663 Colonization Neisseria, 603, 605–6, 610 of oral cavity Alysiella, 833–34 Simonsiella, 833–34 Colony morphology Acidomonas, 179 Actinobacillus actinomycetemcomitans, 845 aerobic phototrophic bacteria, 571 Agrobacterium, 92 Alysiella, 835–37, 837 Azoarcus, 877 Azospirillum, 121–26 Bartonella, 469 Beijerinckia, 155, 157–59 Brucella, 341–44 Burkholderia, 850 Cardiobacterium hominis, 844 Derxia, 753–54, 755–56 Eikenella corrodens, 841–42 Haemophilus aphrophilus, 845 Kingella, 845

900

Index

Leptothrix, 768–70 Leptothrix-Sphaerotilus group, 762 Methylobacter, 278 Methylocaldum, 277 Methylococcus, 275 Methylocystis, 280 Methylomicrobium, 279 Methylomonas, 277 Methylosinus, 280 Phenylobacterium, 253 Simonsiella, 835–37, 837 Sphaerotilus, 773 Xanthobacter, 294–95, 300 Colorless sulfur bacteria. See Thiobacillus Comamonadaceae, 16–17, 723–24, 726, 758 Comamonas, 6, 16, 18, 720, 723–32 applications of, 732 characteristics of, 727 cultivation of, 728 ecology of, 731 genetics of, 729–31 habitat of, 726 history of, 725 identification of, 726–28 isolation of, 726 morphology of, 726 pathogenicity of, 731–32 phylogenetic tree of, 724 phylogeny of, 723 physiology of, 728–29 preservation of, 728 strain of, 728–32 taxonomy of, 723–26 Comamonas acidovorans, 723, 725–27, 850 Comamonas compransoris, 725 Comamonas denitrificans, 723, 726, 728 Comamonas nitrativorans, 723, 726, 728 Comamonas percolans, 723, 725 Comamonas terrigena, 723, 725–26, 728, 731–32 Comamonas testosteroni, 723, 725–32 Compost biofilter, Azoarcus, 875 Conferva ochracea, 758 Conjugation, Neisseria, 626 Contagious abortion, Brucella, 372 Contagious equine metritis, 690 Corroding colonies Eikenella corrodens, 840–45 species capable of, 843–45 Corrosion of agar, 843–45 Eikenella corrodens, 841–42 Thiobacillus, 815 Corynebacterium, 749 Cow manure, Aquaspirillum, 711 Cowdria, 13, 460 Cowdria ruminantium, 460 Cowdriosis. See Heartwater disease Coxiella, 21, 462, 529–41 applications of, 541 cultivation of, 533–34 epidemiology of, 536

genetics of, 535 habitats of, 530 identification of, 531–33 isolation of, 530–31 morphology of, 529 pathogenicity of, 536–41 phylogeny of, 529–30 physiology of, 534–35 Coxiella burnetii, 462, 493, 529–41 Craurococcus, 562, 566 Craurococcus roseus, 163, 562, 565 Crown gall tumors Agrobacterium, 91, 94, 105, 109 biocontrol of, 106–9 Cultivation Acetobacter, 176 Acetobacteraceae, 175 Acidithiobacillus ferrooxidans, 823–24 Acidomonas, 178 Ancalomicrobium, 68–69 Asaia, 180 Azospirillum, 122–23 Beijerinckia, 154 Brucella, 338–41 Comamonas, 728 Coxiella, 533–34 Derxia, 752–53 dimorphic prosthecate bacteria, 83–85 Gluconacetobacter, 181 Gluconobacter, 186 Kozakia, 190 manganese-oxidizing bacteria, 226–28 methanotrophs, 272–74 Methylobacterium, 259–61 Phenylobacterium, 251–52 Prosthecomicrobium, 68–69 Spirillum, 703–6 Thiobacillus, 816–19 Xanthobacter, 299–300 Zymomonas, 202 Cycloclasticus, 20 Cystic fibrosis, 852 Achromobacter, 678, 680 Burkholderia, 848, 849, 850, 851, 856 Cystobacter, 22 Cystobacteraceae, 22 Cysts Beijerinckia, 156–57 Methylobacter, 278, 281–82 Methylocaldum, 275–77 Methylocystis, 282 Methylomonas, 277 Cytochrome a, Beijerinckia, 160 Cytochrome c Alphaproteobacteria, 13 Beijerinckia, 160 Gluconacetobacter, 183 Gluconobacter, 186–87 Paracoccus, 243–44 Zymomonas, 211–12 Cytochrome c oxidase production Methylocystis, 280 Methylosinus, 280 Cytochrome o, Gluconobacter, 186–87

Cytochromes aerobic phototrophic bacteria, 571 Brucella, 367 dimorphic prosthecate bacteria, 74 Paracoccus, 243–44 Zymomonas, 211–12 Cytokines, Neisseria, 616

D Dalapon degradation, Achromobacter/Alcaligenes strains, 685 Dechloromonas, 17, 888 Dechlorosoma, 17 Dechlorosoma suillum, 888 Defensins, 664 Deformin production, Bartonella, 471–72, 477 Dehalogenation, Xanthobacter, 307 Dehalospirillum, 25 Dehalospirillum multivorans, 25 Delftia, 16, 723, 725 Delftia acidovorans, 723, 726 Deltaproteobacteria, 3–29 bdellovibrio group, 23 morphology of, 21 myxobacteria, 22–23 sulfate- and sulfur-reducing bacteria, 23–24 Denitrification Achromobacter/Alcaligenes strains, 680–82 Azoarcus, 885 Chromobacterium, 741 Paracoccus, 238, 240–45 phototrophic alphaproteobacteria, 55 Seliberia, 588 Thiobacillus, 817 Dental plaque, Neisseria, 603 1-Deoxynojirimycin production, Gluconobacter, 189 Dermonecrotic toxin (DNT), Bordetella, 658, 660–61 Derxia, 155, 751–56 applications of, 756 cultivation of, 752–53 habitat of, 751 identification of, 753–54 isolation of, 751–52 morphology of, 753–54, 754–55 motility of, 755 physiology and biochemistry of, 755–56 preservation of, 753 species description of, 754–55 Derxia gummosa, 751–56 Derxia indica, 751 Desulfacinum, 24 Desulfobacter, 23–24 Desulfobulbus, 24 Desulfofaba, 24 Desulfofrigus, 24 Desulfohalobium, 23 Desulfomicrobium, 23 Desulfomonile, 24

Index Desulfonatronum, 23 Desulfonema, 23 Desulforhabdus, 24 Desulfosarcina, 23 Desulfovibrio, 23 Desulfovirga, 24 Desulfurella, 23–24 Desulfuromonas, 23 Desulfuromusa, 23 Di- and Trisaccharide utilization Acidomonas, 179 Asaia, 180 Azospirillum, 123 Kozakia, 190 Zymomonas, 202–3, 208–11 Diagnosis Burkholderia, 852 gonorrhea, 630 of gonorrhea, 604, 606 melioidosis, 852 of meningococcal disease, 604, 631 pertussis, 652–53 Dialysis sac culture, Spirillum, 702 Diazosomes, 885 Dichelobacter, 20 Dichelobacter nodosus, 20 Dihydroxyacetone production, Gluconobacter, 189 3,6-Dihydroxyindoxazene, 741 N,N-Dimethylformamide degradation, Achromobacter/Alcaligenes strains, 686 Dimethylsulfide (DMS) production, Alcaligenes strains, 683 Dimorphic prosthecate bacteria, 72–86 cultivation of, 83–85 distribution of, 75–77 ecology and applications of, 85–86 enrichment and isolation of, 78–81 identification of, 81–83 morphology of, 81–82 rationale for clustering of, 72–75 2,4-Dinitrotoluene degradation, Achromobacter/Alcaligenes strains, 685–86 Disease. See also Animal disease; Human disease; Plant disease agricultural, 651 Agrobacterium, 91–110, 105–9 Bartonella, 467–69, 480–84 Brucella, 381–429 Burkholderia, 852–53 Chromobacterium, 739 Ehrlichia, 514–15 Eikenella corrodens, 843–44 gonorrhea, 629–31 Herbaspirillum, 143 meningitis, 629–32 Neorickettsia, 517 Orientia, 507 pertussis, 650–51 Proteobacteria, 11–12 Rickettsia, 500–504 Rickettsiales, 493 Wolbachia, 516, 555

Disseminated gonococcal infection, 631 D-lysine production, Comamonas, 732 DMS. See Dimethylsulfide production DNA content, Neisseria, 624–28 DNA-DNA hybridization Acetobacter, 176 Acetobacteraceae, 166 Acidomonas, 179 aerobic phototrophic bacteria, 571 ammonia-oxidizing bacteria, 780, 783–85, 789, 793, 795 Ancalomicrobium, 65 Asaia, 180 Azospirillum, 115–16, 123, 125–26, 130 Bartonella, 478 Brucella, 315, 331, 334, 337 Coxiella, 529 dimorphic prosthecate bacteria, 74 Gluconacetobacter, 181 Gluconobacter, 186 Herbaspirillum, 141–42 Kozakia, 190 Methylobacterium, 262 Methylomonas, 269 Phenylobacterium, 254–55 phototrophic alphaproteobacteria, 49–50 Prosthecomicrobium, 65 DNA-rDNA hybridization, Herbaspirillum, 141 DNA-rRNA hybridization Acetobacteraceae, 163, 166 Azospirillum, 115–16, 123 Methylobacterium, 263 Proteobacteria, 5–6 DNT. See Dermonecrotic toxin Dogs Bordetella, 651 Neisseria, 603 Simonsiella, 829, 834 Dolphin habitat, Brucella, 335 Donkeys Alysiella, 832 Burkholderia, 849 Simonsiella, 832 Doxycycline sensitivity Brucella, 429 Coxiella, 537 Ehrlichia, 514 Orientia, 507 Rickettsia, 500, 505–6 Duganella, 888

E Ear habitat Ochrobactrum, 748 Oligella, 691 Ecology aerobic phototrophic bacteria, 580 Agrobacterium, 105 Alphaproteobacteria, 9–15 Alysiella, 837–38

901

Anaplasma, 510 Azoarcus, 886–87 Azonexus, 888 Azospira, 888 Azospirillum, 116–20 Azovibrio, 888 Betaproteobacteria, 15–18 Comamonas, 731 dimorphic prosthecate bacteria, 85–86 Ehrlichia, 513–14 Gammaproteobacteria, 18–21 Herbaspirillum, 143–44 Leptothrix-Sphaerotilus group, 762–64 Neorickettsia, 517 Paracoccus, 237–39 Proteobacteria, 9–25 Rickettsia, 500 Simonsiella, 837–38 Wolbachia, 515–16, 553–55 Ecophysiology, Bordetella, 653–62 Ectothiorhodospiraceae, 18–19 Ehrlicheae, 460–61 Anaplasma, 460–61 Cowdria, 460 Ehrlichia, 460–61 Ehrlichia, 13, 460–61, 493–94, 512–15, 547 disease from, 514–15 ecology of, 513–14 epidemiology of, 514 genetics of, 512–13 habitats of, 512 identification of, 512 isolation of, 512 morphology of, 460 phylogeny of, 512 physiology of, 512 preservation of, 512 taxonomy of, 512 Ehrlichia bovis, 460 Ehrlichia canis, 460, 512–14 Ehrlichia chaffeensis, 460, 510, 512–14 Ehrlichia equi, 460–61 Ehrlichia ewingii, 460, 494, 512, 514 Ehrlichia muris, 460, 512 Ehrlichia phagocytophila, 460–61 Ehrlichia platys, 460 Ehrlichia risticii, 460–61 Ehrlichia ruminantium, 494, 512–15 Ehrlichia sennetsu, 460–61 Ehrlichiaceae, 13 Ehrlichioses Ehrlichia, 514–15 prevention of, 515 Eikenella, 18, 828 Eikenella corrodens, 840–45 antigenic structure of, 842 antimicrobial susceptibility of, 844 biochemical characteristics of, 842 differentiation of, 843 growth characterization of, 841–42 habitat of, 840 identification of, 842 isolation of, 840–42 media for selection of, 840–41

902

Index

microscopic morphology of, 842 pathogenicity of, 842–44 Elephants Alysiella, 832 Simonsiella, 832 Embden-Meyerhof-Parnas pathway aerobic phototrophic bacteria, 577 Coxiella, 534 Paracoccus, 245 Endocarditis Bartonella, 462, 480, 482–84 Coxiella, 537–39 Endoplasmic reticulum, Brucella, 397–400, 407–9 Endosymbiosis methanotrophs, 272 Phyllobacterium, 749 Enterobacter agglomerans, 731 Enterobacteriaceae, 19–20, 727, 843 Brenneria, 19 Buchnera, 19 Erwinia, 19 Escherichia, 19 Pectobacterium, 19 Photorhabdus, 19 Salmonella, 19 Shigella, 19 Xenorhabdus, 19 Yersinia, 19 Entner-Doudoroff pathway aerobic phototrophic bacteria, 576–77 Brucella, 363 Gluconobacter, 188 Paracoccus, 245 Zymomonas, 203 Environmental stress, Brucella, 367–69, 403 Enzymes Agrobacterium, 94–96 Bartonella, 473 Brucella, 362–63 Coxiella, 532 Phenylobacterium, 255 Eperythrozoon ovis, 493 Epidemiology Anaplasma, 510–11 Bartonella, 479–80 Brucella, 372–81 Burkholderia, 851–52 Coxiella, 536 Ehrlichia, 514 Neisseria, 628–30 Orientia, 507 Rickettsia, 500 Epididymitis, Neisseria, 630–31 Epoxide formation, Xanthobacter, 307–8 Epsilonproteobacteria, 3–29 Campylobacteraceae, 24–25 Helicobacter group, 25 morphology of, 24 Equine habitat Burkholderia, 849 Comamonas, 726 Ehrlichia, 460 Taylorella equigenitalis, 689–90

Erwinia, 19 Erythritol pathway, Brucella, 364–67 Erythrobacter, 14, 562, 565, 570–72 Erythrobacter litoralis, 569, 573, 578, 580 Erythrobacter longus, 565, 570–71, 575–77 Erythrocyte parasitism Anaplasma, 508–9, 511 Anaplasmataceae, 493 Bartonella, 472–78, 484 Erythromicrobium, 14, 562, 565, 570–72, 576–77 Erythromicrobium ezovicum, 569–70, 572–73, 578 Erythromicrobium hydrolyticum, 569–71, 575–76, 578 Erythromicrobium ramosum, 570, 573, 575, 578, 580 Erythromonas, 562, 565, 571–72 Erythromonas ursincola, 565, 569, 573, 578 Erythromycin resistance Neisseria, 625 Xanthobacter, 303 Erythromycin sensitivity Coxiella, 537 Xanthobacter, 303 Escherichia, 19, 650 Escherichia coli, 19, 28, 361, 613, 667–68, 864 Estuarine environment aerobic phototrophic bacteria, 566 dimorphic prosthecate bacteria, 76 Methylobacter, 278 Seliberia, 585 Thiobacillus, 815 Ethanesulfonate degradation, Achromobacter/Alcaligenes strains, 684–85 Ethanol fermentation, Zymomonas, 203, 207–9, 212–14 Ethanol oxidation Acetobacter, 176–77 Acetobacteraceae, 163 Acidomonas, 179 Asaia, 179–80 Erythromicrobium, 576 Gluconacetobacter, 180–81, 183 Kozakia, 190 Ethanol tolerance, Zymomonas, 208–9 Eukaryotic cell prokaryotic endosymbionts of, 25–28, 463 proteobacteria associated with, 318–19 Rickettsia, 496 Rickettsiales, 457 Wolbachia, 547 Evolution Bordetella, 650 Simonsiella, 830–31 Exopolysaccharide, Agrobacterium, 97–99 Exospores, Methylosinus, 282 Eye habitat, Ochrobacterum, 748

F False branching Leptothrix discophora, 769 Leptothrix mobilis, 769 Leptothrix-Sphaerotilus group, 759 Sphaerotilus, 773 Fatty acid utilization aerobic phototrophic bacteria, 576 phototrophic alphaproteobacteria, 55–56 Fatty acids Acetobacteraceae, 163 Achromobacter, 676 Acidomonas, 179 aerobic phototrophic bacteria, 571 Alcaligenes, 676 Alphaproteobacteria, 13 Alysiella, 837 Bartonella, 470–71 Brucella, 316, 320, 350–51 Coxiella, 532 Kerstersia, 676 methanotrophs, 271–72 Methylobacter, 278 Methylocaldum, 277 Methylococcus, 275 Methylocystis, 280 Methylomicrobium, 279 Methylomonas, 277–78 Methylosinus, 281 Methylosphaera, 279 nomenclature of, 275 Oligella, 690 Paracoccus, 240–43 Pelistega, 690 Pigmentiphaga, 676 Proteobacteria, 7 Simonsiella, 837 Taylorella, 690 Xanthobacter, 303–4 Feces Achromobacter/Alcaligenes strains, 687 Alcaligenes, 677 Comamonas, 726 Ochrobactrum, 748 Feline habitat Bartonella, 462, 469, 479 Coxiella, 536 Simonsiella, 829, 834 Ferric hydroxide, LeptothrixSphaerotilus group, 759 Ferromanganese oxides, LeptothrixSphaerotilus group, 759 Filament Alysiella, 833–35 Simonsiella, 828, 833–35 Filamentous bacteria, 758–75 Alysiella, 828–38 Simonsiella, 828–38 Filamentous hemagglutinin, 663, 667 Bordetella, 656–57 Fimbriae, 667 Bordetella, 657, 665 Fish habitat Coxiella, 530

Index dimorphic prosthecate bacteria, 77 Fission, dimorphic prosthecate bacteria, 73 Flagella Acidomonas, 179 aerobic phototrophic bacteria, 571–72 Agrobacterium, 96 Azospirillum, 123, 125, 127 Bartonella, 462, 467, 470–71, 474–75, 477 bdellovibrio group, 23 Beijerinckia, 156–58 Burkholderia, 856 dimorphic prosthecate bacteria, 72–73, 81, 83 Epsilonproteobacteria, 24 Gluconacetobacter, 180 Gluconobacter, 185 Methylobacter, 278 Methylobacterium, 261 Methylomicrobium, 278 Methylomonas, 277 Proteobacteria, 3 Xanthobacter, 296, 301 Flavobacterium capsulatum, 742 Flour, Acetobacteraceae, 174 Flowers Acetobacter, 176 Acetobacteraceae, 172–73 Asaia, 179–80 Gluconacetobacter, 181 Gluconobacter, 186 Fluorobenzoate degradation, Achromobacter/Alcaligenes strains, 685 Fluoroquinolone sensitivity Brucella, 428 Coxiella, 537 Food, Janthinobacterium, 742 Formaldehyde production, methanotrophs, 266 Formate production, phototrophic alphaproteobacteria, 55 Formate utilization, phototrophic alphaproteobacteria, 56 Francisella, 20, 461 Francisella tularensis, 20 Frateuria, 21 Freshwater environment aerobic phototrophic bacteria, 562, 566 ammonia-oxidizing bacteria, 783–86, 804 Ancalomicrobium, 65–66 Aquaspirillum, 711 Azospirillum, 122 bdellovibrio group, 23 dimorphic prosthecate bacteria, 76, 83 Leptothrix, 764–65 Leptothrix cholodnii, 768 Leptothrix discophora, 769 Leptothrix lopholea, 768 Leptothrix mobilis, 769 Leptothrix pseudoochracea, 768 methanotrophs, 270–72 Methylobacterium, 258–59

nitrite-oxidizing bacteria, 863–66 phototrophic alphaproteobacteria, 44–45 phototrophic betaproteobacteria, 596 Prosthecomicrobium, 65–66 Seliberia, 585–86 Skermanella, 126 Sphaerotilus, 772 Spirillum, 701 Thiobacillus, 813, 815 Thiovulum, 25 Xanthobacter, 295 Fruit tree habitat, Agrobacterium, 92 Fruits Acetobacter, 176 Acetobacteraceae, 172–73 Gluconacetobacter, 181 Gluconobacter, 186

G Gallionella ferruginea, 17, 770 Gammaproteobacteria, 3–29 Aeromonadaceae, 19–20 Alteromonadaceae, 19–20 ammonia-oxidizing bacteria, 786 Cardiobacteriaceae, 20 Chromatiaceae, 18–19 Ectothiorhodospiraceae, 18–19 Enterobacteriaceae, 19–20 Halomonadaceae, 20 Legionellaceae, 21 Methylococcaceae, 20 Moraxellaceae, 20 orders and families of, 18 Pasteurellaceae, 19–20 Pseudomonadaceae, 20–21 Succinivibrionaceae, 19–20 sulfur oxidizing bacteria of, 812–14, 821–24 Vibrionaceae, 19–20 Gas vesicles Methylobacter, 278 Methylosphaera, 279 Prosthecomicrobium, 70 Gastrointestinal tract Eikenella corrodens, 840 Enterobacteriaceae, 19 G+C content Acetobacter, 177 Achromobacter, 676 aerobic phototrophic bacteria, 571 Alcaligenes, 676, 684 Alcaligenes defragrans, 677 Alcaligenes denitrificans, 689 Alcaligenes faecalis, 677, 683, 687–88 Alcaligenes faecalis S-6, 680 Alysiella, 832, 837 ammonia-oxidizing bacteria, 784–86, 795 Anaplasma, 509 Aquaspirillum, 710–11, 713 Azoarcus, 877, 879–85 Azonexus, 889 Azospira, 889 Azospirillum, 116

903

Azovibrio, 889 Bartonella, 478 Beijerinckia, 158–59 Brucella, 320, 354, 372 Chromobacterium, 738 Comamonas, 728 Coxiella, 535 Derxia, 755 dimorphic prosthecate bacteria, 74 Eikenella corrodens, 840 Gluconacetobacter, 183 Herbaspirillum, 142 Janthinobacterium, 741 Kerstersia, 676 Leptothrix, 766 Leptothrix-Sphaerotilus group, 761 Methylobacter, 278 Methylocaldum, 277 Methylococcus, 275 Methylocystis, 280 Methylomicrobium, 279 Methylomonas, 277 Methylosinus, 281 Methylosphaera, 279 Oligella, 675, 690–91 Paracoccus, 234, 240–43 Pelistega, 675, 690 Phenylobacterium, 254 phototrophic alphaproteobacteria, 50 Pigmentiphaga, 676 proteobacteria, 28 Rickettsia, 498 Simonsiella, 832, 837 Spirillum, 704–5 Taylorella, 675, 689–90 Xanthobacter, 290 Gelatin hydrolysis, phototrophic betaproteobacteria, 599 Gemmobacter, 14 Gene expression, Brucella, 371–72 Gene structure, Bartonella, 478–79 Gene transfer, Zymomonas, 212 Genetic engineering, Agrobacterium, 109–10 Genetic exchange Bartonella, 479 Brucella, 372 Genetics Acetobacter, 177–78 Acidomonas, 179 aerobic phototrophic bacteria, 578–80 Agrobacterium, 96–105 Anaplasma, 509–10 Azoarcus, 886 Azonexus, 890 Azospira, 890 Azospirillum, 126–32 Azovibrio, 890 Bartonella, 478–79 Brucella, 327–28, 330, 332–33, 368–72 clonal and reticulate, 332–33 Comamonas, 729–31 Coxiella, 535 Ehrlichia, 512–13

904

Index

Gluconacetobacter, 184–85 Gluconobacter, 187–88 methanotrophs, 281–84 Neisseria, 624–28 Neorickettsia, 516–17 Orientia, 506–7 Rickettsia, 498–500 selective silencing of, 650 Xanthobacter, 306–7 Zymomonas, 212–14 Genitourinary tract Eikenella corrodens, 840 Ochrobactrum, 748 Oligella, 691 Genome Bartonella, 478 Brucella, 321–24, 368 Coxiella, 535 Rickettsia, 498 Wolbachia, 552 Gentamycin sensitivity, Methylobacterium, 262 Geobacter, 24 Glanders, 848, 849, 851–52, 853, 856 Gluconacetobacter, 13, 163–66, 180–85 applications of, 185 characteristics of, 168, 181, 186–87 classification of, 167, 180, 182 genetics of, 184–85 habitats of, 181 identification of, 181 isolation, cultivation and preservation of, 181 taxonomy of, 180–81 Gluconacetobacter azotocaptans, 173, 181, 184 Gluconacetobacter diazotrophicus, 119, 126, 173, 180–81, 184–85 Gluconacetobacter entanii, 172, 175, 181, 183 Gluconacetobacter europaeus, 175, 180–81, 183, 185 Gluconacetobacter hansenii, 173, 180–81, 183 Gluconacetobacter intermedius, 174, 181, 183–85 Gluconacetobacter johannae, 173, 181, 184 Gluconacetobacter liquefaciens, 169, 180–81 Gluconacetobacter oboediens, 181, 183 Gluconacetobacter sacchari, 173 Gluconacetobacter xylinus, 169, 171–74, 180, 183–84 Gluconacetobacter xylinus subsp. sucrofermentans, 183 Gluconate production Acidomonas, 179 Gluconacetobacter, 185 Gluconobacter, 188 Kozakia, 190 Gluconeogenesis, Coxiella, 534 Gluconobacter, 13, 151, 163–66, 168–70, 174, 185–89 applications of, 188–89 characteristics of, 168 classification of, 167

genetics of, 187–88 habitats of, 186 identification of, 186 isolation and cultivation of, 186 taxonomy of, 185–86 Gluconobacter asaii, 186 Gluconobacter cerinus, 186 Gluconobacter frateurii, 172–73, 186 Gluconobacter oxydans, 166, 169, 172–74, 185–86, 188–89 Glucose utilization Acidomonas, 179 Gluconacetobacter, 181 Gluconobacter, 186–87 Paracoccus, 245 Glutamate dehydrogenase, phototrophic betaproteobacteria, 598 Glycerol utilization Acidomonas, 179 Brucella, 363–64 Glycolytic pathway, Brucella, 364 Glycoproteins, Ehrlichia, 513 Glycosphingolipids, Sphingomonadaceae, 14 Goat habitat. See Caprine habitat Gold solubilization, Chromobacterium, 741 Gonococcal arthritis, Neisseria, 631 Gonococcus. See Neisseria gonorrhoeae Gonorrhea, 603, 630–31 antibiotic treatment of, 632 complications of, 630–31 diagnosis of, 604, 606, 630 disease incidence of, 629 prevention of, 631 Grahamella, 493. See also Bartonella Grape habitats Acetobacteraceae, 168–69 Agrobacterium, 91–92, 105–6, 109 Gluconobacter, 186 Grass roots, Azoarcus, 873, 875 Guinea pig habitat. See Rodent habitat

H Haemobartonella, 473 Haemophilus, 19, 841–42, 845 Haemophilus aphrophilus, 844–45 Haemophilus equigenitalis, 689 Haemophilus influenzae, 19, 28 Haemophilus parainfluenzae, 625 Haemophilus paraphrophilus, 844 Hairy root disease, Agrobacterium, 91, 105–6 Halomonadaceae, 20 Carnimonas, 20 Chromohalobacter, 20 Halomonas, 20 Zymobacter, 20 Halomonas, 20 Halothiobacillus, 18 Halothiobacillus neapolitanus, 816 Hazardous waste degradation, Xanthobacter, 308–9 Heartwater disease, Ehrlichia, 515

Heat shock proteins Agrobacterium, 101 Brucella, 403–4 Heavy metals detection of, Spirillum, 707–8 nitrite-oxidizing bacteria, 869 resistance to Achromobacter/Alcaligenes strains, 686 Comamonas, 731 Helical bacteria Aquaspirillum, 710–21 Spirillum, 701–8 Helicobacter bovis, 25 Helicobacter group, 25 Helicobacter, 25 Nautilia, 25 Thiomicrospira denitrificans, 25 Thiovulum, 25 Wolinella, 25 Helicobacter pylori, 24–25 Helicobacter suis, 25 Heme, Neisseria binding of, 615–16 Hemobartonella felis, 493 Hemobartonella muris, 493 Hemoglobin, Neisseria binding of, 615–16 Hemolysin production, Bartonella, 471–72 Hemolysis Bordetella, 660 Burkholderia, 854 Simonsiella, 835 Heparan sulfate proteoglycan (HSPG) receptors, 611 Herbaspirillum, 17, 141–47 applications of, 147 characteristics of, 145 habitats and ecology of, 143–44 history of, 141 identification of, 145–46 isolation of, 144 morphology of, 145 phylogenetic tree of, 142 preservation of, 144–45 taxonomy of, 141–43 Herbaspirillum chlorophenolicum, 141, 143–45 Herbaspirillum frisingense, 141–44, 146–47 Herbaspirillum lusitanum, 141, 143–44, 146 Herbaspirillum rubrisubalbicans, 141, 143–44, 146 Herbaspirillum seropedicae, 141–44, 146–47 Herbicide degradation, Phenylobacterium, 250 Hippea, 23 Hippopotami Alysiella, 832 Simonsiella, 832 Hirschia, 14 Holdfasts Leptothrix discophora, 768 Leptothrix lopholea, 768 Leptothrix-Sphaerotilus group, 759, 762 Sphaerotilus, 773

Index Holospora Caedibacter, 462 Holospora, 462 Holospora, 13, 462 Holospora obtusa, 462 Honey habitat, Zymomonas, 201 Hopanoids, Zymomonas, 208–9 Horizontal transmission Bordetella, 650 Simonsiella, 828 Horse habitat. See Equine habitat Horse manure, Aquaspirillum, 711 Hospital environment Burkholderia, 850, 851 Comamonas, 726 Methylobacterium, 258–59 Ochrobactrum, 748 Hot spring environment aerobic phototrophic bacteria, 565 phototrophic alphaproteobacteria, 45 Thiobacillus, 815 Human disease Alphaproteobacteria, 14–15 Anaplasma, 511 Anaplasmataceae, 493 Bartonella, 462, 467–69, 479 Betaproteobacteria, 16 Bilophila, 23 Brucella, 315, 335–38, 372–429 Campylobacteraceae, 24–25 Cardiobacteriaceae, 20 Chlamydia, 463 Coxiella, 531, 536 Ehrlichia, 460–61, 514–15 Enterobacteriaceae, 19 Francisella, 20 Gammaproteobacteria, 18 Legionella, 21 Neisseriaceae, 18 Orientia, 459–60 Pasteurellaceae, 19 Proteobacteria, 11 Rickettsia, 456–60 Human granulocytic ehrlichiosis, Ehrlichia, 460–61 Human habitats Achromobacter, 678 Alcaligenes, 677 Alysiella, 832, 833–34 Bordetella, 650–51 Eikenella corrodens, 840 Neisseria, 603 Ochrobactrum, 748 Simonsiella, 829, 832, 833–34 Human monocytotropic ehrlichiosis, Ehrlichia, 460, 514 Hydrazine degradation, Achromobacter/Alcaligenes strains, 686 Hydrogen oxidation Paracoccus, 238 Xanthobacter, 290 Hydrogen production, phototrophic alphaproteobacteria, 55, 58–59 Hydrogenophaga, 16, 723 Hydrogenophaga flava, 16 Hydrogenophilus, 17

Hydrogenovibrio, 20 Hydrophilic pathway, Brucella, 359–60 Hydrophobic pathway, Brucella, 360 Hydrothermal environment aerobic phototrophic bacteria, 566 dimorphic prosthecate bacteria, 75–76 Hydrothermal vents, Thiobacillus, 815 3-(3-Hydroxyphenyl)-propionate degradation, Comamonas, 730 Hydroxypyruvate reductase, Paracoccus, 235 Hyperinduction, Azoarcus, 885 Hypersaline environment ammonia-oxidizing bacteria, 804 phototrophic alphaproteobacteria, 45 Hyphomicrobiaceae, 15 Hyphomicrobium, 15 Pedomicrobium, 15 Rhodomicrobium, 15 Xanthobacter, 290, 294 Hyphomicrobium, 15, 65, 72–86, 225 Hyphomicrobium aestuarii, 75, 80 Hyphomicrobium facilis, 80 Hyphomicrobium vulgare, 80 Hyphomicrobium zavarzinii, 80 Hyphomonas, 14, 72–86 Hyphomonas hirschiana, 74 Hyphomonas neptunium, 85 Hyphomonas polymorpha, 75, 77 Hypocreales, 885

I Identification Acetobacter, 176 Achromobacter/Alcaligenes strains, 686–87 Acidomonas, 178–79 aerobic phototrophic bacteria, 568–71 Agrobacterium, 92–94 Alysiella, 835–37 Anaplasma, 508–9 Ancalomicrobium, 69–70 Aquaspirillum, 719–21 Asaia, 180 Azoarcus, 877–85 Azonexus, 888–89 Azospira, 888–89 Azospirillum, 123–26 Azovibrio, 888–89 Bartonella, 469–71 Beijerinckia, 154–57 Brucella, 325–26, 341–48 Burkholderia, 850–51 Chromobacterium, 740 Comamonas, 726–28 Coxiella, 531–33 Derxia, 753–54 dimorphic prosthecate bacteria, 81–83 Ehrlichia, 512 Eikenella corrodens, 842

905

Gluconacetobacter, 181 Gluconobacter, 186 Herbaspirillum, 145–46 Janthinobacterium, 742–43 Kozakia, 190 Leptothrix, 766–69 Leptothrix-Sphaerotilus group, 763–64 manganese-oxidizing bacteria, 228–29 methanotrophs, 274–81 Methylobacterium, 261–63 Methylococcaceae, 275–79 Methylocystaceae, 279–81 Neisseria, 604–6 Ochrobactrum, 748–49 Orientia, 506 Phenylobacterium, 252–56 phototrophic alphaproteobacteria, 49–54 phototrophic betaproteobacteria, 597–98 Phyllobacterium, 748–49 Prosthecomicrobium, 69–70 Rickettsia, 496 Seliberia, 588 Simonsiella, 835–37 Sphaerotilus, 773–74 Spirillum, 703 Thiobacillus, 820–21 Wolbachia, 551–52 Xanthobacter, 300–306 Zymomonas, 202–3 Ideonella, 758–59 Iminodisuccinate degradation, Achromobacter/Alcaligenes strains, 686 Immunity Bartonella, 484–85 to pertussis, 664–66, 667 Incidence gonorrhea, 629 meningitis, 629–30 Inclusion bodies, Xanthobacter, 301 Indole degradation, Achromobacter/Alcaligenes strains, 686 Industrial processes, LeptothrixSphaerotilus group, 763 Innate immune defenses, in respiratory tract, 662–64 Insertion sequence Brucella, 370–71 Coxiella, 535 Gluconacetobacter, 184 Interleukins, Brucella, 414–15 Intracellular state, Bordetella, 667–68 Intravenous tubing, Comamonas, 726 Iodobacter, 737–38 Iodobacter fluviatilis, 742 Iron chelation, Brucella, 405–6 Iron (Fe2+) oxidation Acidithiobacillus ferrooxidans, 823 Leptothrix, 770–71 Leptothrix ochracea, 766–67 Sphaerotilus, 773–74

906

Index

Iron seeps and springs, Leptothrix, 764–65 Iron uptake, Brucella, 362–63 Iron-binding proteins, Neisseria, 613–16 Isoenzyme typing, Neisseria, 628 Isolation Acetobacter, 176 Acetobacteraceae, 175 Acidithiobacillus ferrooxidans, 823 Acidithiobacillus thiooxidans, 824 Acidomonas, 178 aerobic phototrophic bacteria, 566–68 Agrobacterium, 92 Alysiella, 834–35 ammonia-oxidizing bacteria, 778–79 Anaplasma, 508 Ancalomicrobium, 66–68 Aquaspirillum, 711–19 Asaia, 180 Azoarcus, 876–77 Azonexus, 888 Azospira, 888 Azospirillum, 120–22 Azovibrio, 888 Bartonella, 469 Beijerinckia, 152–54 Bordetella, 651–53 Brucella, 337–41 Burkholderia, 850 Chromobacterium, 739–40 Comamonas, 726 Coxiella, 530–31 Derxia, 751–52 dimorphic prosthecate bacteria, 78–81 Ehrlichia, 512 Eikenella corrodens, 840–42 Gluconacetobacter, 181 Gluconobacter, 186 Herbaspirillum, 144 Janthinobacterium, 742 Kozakia, 190 Leptothrix, 765–66 manganese-oxidizing bacteria, 225–28 methanotrophs, 272–74 Methylobacterium, 259–61 Neisseria, 603–4 nitrite-oxidizing bacteria, 865–66 Ochrobactrum, 748 Orientia, 506 Paracoccus, 237–40 Phenylobacterium, 250–51 phototrophic alphaproteobacteria, 45–49 phototrophic betaproteobacteria, 596–97 Phyllobacterium, 748 Prosthecomicrobium, 66–68 Rickettsia, 496 Seliberia, 585–88 Simonsiella, 834–35 Sphaerotilus, 772–73 Spirillum, 701–3 Thiobacillus, 816–19

Wolbachia, 551 Xanthobacter, 297–99 Zymomonas, 201–2 Isoprene degradation, Alcaligenes strains, 682

J Janthinobacterium, 17, 720, 737, 741–43 applications of, 743 differentiating characteristics of, 738, 743 enrichment of, 742 habitat of, 741–42 identification of, 742–43 isolation of, 742 morphology of, 741 motility of, 741 preservation of, 743 Janthinobacterium lividum, 737, 740, 741–43, 751

K Kallar grass Azoarcus, 875, 887 Azospira, 888 Azovibrio, 888 Kanamycin sensitivity, Methylobacterium, 262 Kennel cough, 651 Kerstersia, 675–76, 678–89 physiology of, 679 Kerstersia gyiorum, 689 Ketogluconate production Gluconobacter, 185, 188 Kozakia, 190 Ketogulonicigenium, 14 Kidneys, Comamonas, 726 Kingella, 18, 828, 844–45 Kingella denitrificans, 845 Kingella indologenes, 845 Kingella kingae, 845 Klebsiella, 747, 749 Klebsiella oxytoca, 854 Klebsiella pneumoniae, 146 Knallgas bacteria, Xanthobacter, 290 Kombucha Acetobacteraceae, 174 Gluconacetobacter, 181 Kozakia, 163–66, 189–90 characteristics of, 168, 190 classification of, 167 habitats of, 190 identification of, 190 isolation and cultivation of, 190 taxonomy of, 189–90 Kozakia baliensis, 174, 180, 190 Krebs’ cycle, Paracoccus, 245

L Lactate oxidation, Acetobacter, 175–76 Lactate production, Beijerinckia, 158

Lactobacillus acidophilus, 607 Lactobacillus viridescens, 174 Lactoferrin, 613–14, 663–64 Neisseria, 614–15 Lantadene degradation, Alcaligenes strains, 682 Lawsonia, 23 Lawsonia intracellularis, 23 Leaf nodules, Phyllobacterium, 747, 749 Legionella, 21, 329 Legionella pneumophila, 21, 330, 499, 529 Legionellaceae, 21 Coxiella, 21 Legionella, 21 Rickettsiella, 21 Leptothrix, 16, 225, 764–72, 774–75 characteristics of, 766 habitat of, 764–65 identification of, 766–69 isolation of, 765–66 metal oxidation by, 770–71 physiology of, 770–72 sheath structure and composition in, 771–72 Leptothrix cholodnii, 759, 762, 765–66, 768–69 Leptothrix cholodnii SP-6, 771–72 Leptothrix discophora, 229–30, 593, 675, 762, 766, 768–70 Leptothrix discophora SS-1, 764, 770–71 Leptothrix lopholea, 759, 762, 766, 768 Leptothrix mobilis, 766, 769–70 preservation of, 769 Leptothrix ochracea, 758, 761–62, 766–67, 773 Leptothrix pseudoochracea, 766–68 Leptothrix-Sphaerotilus group, 758–75 common traits of, 759–61 distinguishing traits of, 761 ecology of, 762–64 morphology of, 761 phenotypic characteristics of, 759–62 phylogeny of, 758–62 physiology of, 761 reference strains of, 762 studying distribution and abundance of, 763–64 taxonomy of, 758–62 Leucothrix, 20 Lice habitat Bartonella, 479 Rickettsia, 500 Ligase chain reaction, for gonorrhea diagnosis, 606 Lincomycin sensitivity, Coxiella, 537 Lipid A Alphaproteobacteria, 7 Brucella, 330, 345, 352–56, 359–61 Coxiella, 532 Legionella, 330 Lipid-modified azurin, Neisseria, 613 Lipids, Brucella, 346, 349–51, 360–61

Index Lipoid bodies, Beijerinckia, 155–58, 160 Lipooligosaccharide Brucella, 353–54 Neisseria, 617–22 antigenic variation of, 617–19, 617–21 biosynthesis of, 620–21 expression of, 618 sialylation of, 618–19 structure of, 618, 619 Lipopolysaccharide Bartonella, 471 Bordetella, 661–62 Brucella, 316, 327–28, 331, 344–45, 349, 351–56, 358–59, 372, 391, 404–5, 410–11, 417–18, 422 Coxiella, 531–32 Phenylobacterium, 254–55 phototrophic alphaproteobacteria, 50 Xanthobacter, 302 Lipoprotein, Rickettsia, 500, 505 Lipoquinone Methylomicrobium, 279 Methylomonas, 278 Lithotrophy, nitrite-oxidizing bacteria, 869 Lophomonas, 723 Lophomonas alcaligenes, 723 LOS. See Lipooligosaccharide LPS receptors, 663 LPS-binding protein (LBP), 663 Lymphocytosis, Bordetella, 659 Lysosome inhibition, Brucella, 404– 5

M Macrolide sensitivity, dimorphic prosthecate bacteria, 74 Macrophages, Brucella, 388, 391–94, 398–401, 405–6, 409–15 Magnetospirillum magnetotacticum, 41 Maize habitat Azospirillum, 118–20, 133 Chromobacterium, 738–39 Herbaspirillum, 143 Malleobactin, Burkholderia, 854 Malonomonas rubra, 23–24 Malta fever, Brucella, 372 Manganese (Mn2+) oxidation Leptothrix, 766–71 Leptothrix-Sphaerotilus group, 759, 761–62 Manganese chemistry, 223–24 Manganese-oxidizing bacteria, 222–30 applications of, 229 controversy and perspectives of, 229–30 habitats of, 222–25 Hyphomicrobium, 225 identification of, 228–29 isolation and enrichment of, 225–28 Leptothrix, 225

Manganese-oxidizing factor, 770–71, 772 Mannheimia, 19 Marine environment aerobic phototrophic bacteria, 562, 566 Alcaligenes strains, 682–83 ammonia-oxidizing bacteria, 802–4 Ancalomicrobium, 65–66 bdellovibrio group, 23 dimorphic prosthecate bacteria, 76 Gammaproteobacteria, 19–20 nitrite-oxidizing bacteria, 863, 866 phototrophic alphaproteobacteria, 44–45 Prosthecomicrobium, 65–66 Psychrobacter, 20 Thiobacillus, 813, 815 Thiovulum, 25 Vibrionaceae, 19 Xanthobacter, 295 Marine mammal habitat, Brucella, 335 Meat, fermented, Acetobacteraceae, 175 Media aerobic phototrophic bacteria, 568 Agrobacterium, 93 Alcaligenes, 677 ammonia-oxidizing bacteria, 779–80 Ancalomicrobium, 68–69 Azospirillum, 120–22 Brucella, 339–43 dimorphic prosthecate bacteria, 83–85 Eikenella corrodens, 840–41 Herbaspirillum, 145 methanotrophs, 272–74 nitrite-oxidizing bacteria, 866 Paracoccus, 239–40 phototrophic alphaproteobacteria, 46–49 Prosthecomicrobium, 68–69 Spirillum, 702–3 Xanthobacter, 299 Melioidosis, 848, 850, 852, 855–56 Melittangium, 22 Menaquinones Deltaproteobacteria, 7–8, 21 Epsilonproteobacteria, 7–8, 24 Meningitis, 631–32. See also Meningococcal disease Meningococcal disease, 603, 631–32 antibiotic treatment of, 632 complications of, 631–32 diagnosis of, 604, 631 disease incidence of, 629–30 prevention of, 632 Meningococcus. See Neisseria meningitidis Mesorhizobium, 14–15 Metabolic engineering, Zymomonas, 212–14 Metabolism Bartonella, 471 Brucella, 349, 361–68

907

methanotrophs, 281 Neisseria, 607–8 Proteobacteria, 4–5 Zymomonas, 203–7 Metal working fluids, Comamonas, 726 Metallogenium, 225, 230 Methane monooxygenase methanotrophs, 266, 272–73, 282–84 Methylobacter, 278 Methylomicrobium, 279 Methylosphaera, 279 particulate, 282–83 soluble, 283 Methane oxidation methanotrophs, 266, 281, 282 Methylobacter, 278 Methylocystis, 280 Methylomicrobium, 279 Methylosphaera, 279 Methanococcus, 703 Methanol utilization Acidomonas, 179 dimorphic prosthecate bacteria, 83–84, 86 methanotrophs, 266, 281 Methylobacterium, 257–60 Methylocystis, 280 Methylomicrobium, 279 Methylosphaera, 279 phototrophic alphaproteobacteria, 56 Methanol yeast process, Acetobacteraceae, 174 Methanotrophs, 266–84. See also Methylococcaceae; Methylocystaceae applications of, 284 characteristics of, 267 habitats of, 270–72 detection, 271–72 endosymbionts, 272 identification of, 274–81 isolation and cultivation of, 272–74 phylogenetic tree of, 267–68 phylogeny of, 266–68 physiology and genetics of, 281–84 preservation of, 281 taxonomy of, 266, 268–70 types of, 266–68 Methylarcula, 14 Methylhydrazine degradation, Achromobacter/Alcaligenes strains, 686 Methylobacillus, 17 Methylobacter, 20, 268–69 identification of, 278–79 taxonomy of, 269 Methylobacter alcaliphilus, 269, 278 Methylobacter luteus, 269, 278 Methylobacter marinus, 269, 278 Methylobacter psychrophilus, 269, 271, 273, 278 Methylobacter whittenburyi, 269, 278 Methylobacterium, 15, 257–63 applications of, 263

908

Index

habitats of, 258–59 isolation and cultivation of, 259–61 morphology of, 261 preservation of, 263 taxonomy of, 257–58 Methylobacterium aminovorans, 258 Methylobacterium chloromethanicum, 258 Methylobacterium dichloromethanicum, 258 Methylobacterium ethanolicum, 258 Methylobacterium extorquens, 258 Methylobacterium fujisawaense, 258 Methylobacterium hypolimneticum, 258 Methylobacterium mesophilicum, 258 Methylobacterium organophilum, 258–59, 261 Methylobacterium rhodesianum, 258 Methylobacterium rhodinum, 258, 571 Methylobacterium thiocyanatum, 258 Methylobacterium zatmanii, 258 Methylocaldum, 266 identification of, 275–77 taxonomy of, 268 Methylocaldum gracile, 268, 277 Methylocaldum szegediense, 268, 277 Methylocaldum tepidum, 268, 277 Methylocapsa, 15 Methylococcaceae, 20, 266–84 characteristics of, 276 identification of, 275–79 Methylobacter, 20, 266 Methylocaldum, 266 Methylococcus, 20, 266 Methylomicrobium, 266 Methylomonas, 20, 266 Methylosphaera, 266 taxonomy of, 268–69 Methylococcus, 20, 266 identification of, 275 taxonomy of, 268 Methylococcus bovis. See Methylobacter luteus Methylococcus capsulatus, 268, 275, 283 Methylococcus chroococcus. See Methylobacter whittenburyi Methylococcus thermophilus, 268, 275 Methylococcus vinelandii. See Methylobacter whittenburyi Methylocystaceae, 266–84 identification of, 279–81 Methylocystis, 266 Methylosinus, 266 taxonomy of, 269–70 Methylocystis, 15, 266 identification of, 279–80 taxonomy of, 269–70 Methylocystis echinoides, 270, 280 Methylocystis minimus, 269–70 Methylocystis parvus, 269–70, 279–80, 282 Methylomicrobium, 266, 269 identification of, 278–79

taxonomy of, 269 Methylomicrobium agile, 269, 279 Methylomicrobium album, 269, 283 Methylomicrobium pelagicum, 269, 271, 279 Methylomonas, 20, 266 identification of, 277–78 taxonomy of, 268–69 Methylomonas aurantiaca, 277 Methylomonas flagellata, 269 Methylomonas fodinarum, 277 Methylomonas margaritae, 269 Methylomonas methanica, 268–69, 277 Methylomonas methaninitrificans, 269 Methylomonas methanooxidans, 269 Methylomonas pelagica. See Methylomicrobium Methylophaga, 20, 266 Methylophilus, 17, 269 Methylosinus, 266, 269 identification of, 280–81 taxonomy of, 270 Methylosinus sporium, 270, 280–81 Methylosinus trichosporium, 270, 280–84 Methylosphaera, 266 identification of, 279 taxonomy of, 269 Methylosphaera hansonii, 269, 271–73, 279, 283–84 Methylovibrio soehngenii, 270 Methylovorus, 17 Mice habitat. See Rodent habitat Micrococcus denitrificans. See Paracoccus denitrificans Micrococcus radiodurans, 688 Miglitol production, Gluconobacter, 189 Milk habitat, Comamonas, 726 Mitochondria Alphaproteobacteria, 3, 7, 13, 25–28 as prokaryotic endosymbiont, 463–64 Rickettsia, 495 MOF. See Manganese-oxidizing factor Molybdenum requirement, Beijerinckia, 160 Monkeys Alysiella, 832 Chromobacterium, 739 Simonsiella, 832 Monoclonal antibody Neisseria, 612, 618–19 nitrite-oxidizing bacteria, 863–65 Monosaccharide utilization Acidomonas, 179 aerobic phototrophic bacteria, 576 Asaia, 180 Azospirillum, 115, 123, 127 Brucella, 362–64, 367 Gluconacetobacter, 181 Gluconobacter, 186–87 Herbaspirillum, 145–46 Kozakia, 190 Paracoccus, 240–43

phototrophic alphaproteobacteria, 55–56 Xanthobacter, 290–91, 305 Zymomonas, 202–3, 208–11, 213 Moraxella, 20, 828, 835, 845 Moraxellaceae, 20 Acinetobacter, 20 Moraxella, 20 Psychrobacter, 20 Morphology aerobic phototrophic bacteria, 571 Agrobacterium, 94 Alphaproteobacteria, 9–15 Alysiella, 828, 830, 834–36 ammonia-oxidizing bacteria, 780, 783–86 Anaplasmataceae, 493 Ancalomicrobium, 65 Aquaspirillum, 719 Azoarcus, 877–85 Azonexus, 889 Azospira, 889 Azospirillum, 115, 121 Azovibrio, 889 Bartonella, 467, 470 bdellovibrio group, 23 Beijerinckia, 151–52, 154–59 Betaproteobacteria, 15–18 Brucella, 341–44, 349–61 Cardiobacterium hominis, 844 Chlamydiae, 463 Chromobacterium, 738 Comamonas, 726 Coxiella, 529 Deltaproteobacteria, 21 Derxia, 754–55 dimorphic prosthecate bacteria, 81–82 Ehrlichia, 460 Eikenella corrodens, 842 Epsilonproteobacteria, 24 Gammaproteobacteria, 18–21 Haemophilus paraphrophilus, 844 Herbaspirillum, 145 Janthinobacterium, 741 Leptothrix, 767–70 Leptothrix-Sphaerotilus group, 759, 761 Methylobacter, 278 Methylobacterium, 261 Methylocaldum, 275 Methylocystis, 279–80 Methylomicrobium, 278 Methylomonas, 277 Methylosinus, 280 Methylosphaera, 279 Myxobacteria, 22 Neisseria, 605–6 Paracoccus, 233–34 Phenylobacterium, 252–53 Prosthecomicrobium, 65, 70 Proteobacteria, 3, 9–25 Rhodocyclus, 597 Rickettsia, 457–59 Rubrivivax, 596 Seliberia, 585, 587 Simonsiella, 828, 830, 833–36 Skermanella, 126 Sphaerotilus, 773–74

Index Spirillum, 703 Wolbachia, 515, 551 Xanthobacter, 290, 294–96, 300–301 Mosquito habitat, Wolbachia, 547, 550–51 Motility Acidomonas, 179 Agrobacterium, 96 Alysiella, 833–35, 833–37 Aquaspirillum, 719 Azoarcus, 877 Azospirillum, 121, 123, 125 Bartonella, 471, 474 Beijerinckia, 156–59 Bordetella, 650 Burkholderia, 850–51, 856 Chromobacterium, 738 Deltaproteobacteria, 21 Derxia, 755 dimorphic prosthecate bacteria, 72–73 Eikenella corrodens, 841 Epsilonproteobacteria, 24 Gluconacetobacter, 180 Gluconobacter, 185 Herbaspirillum, 145 Janthinobacterium, 741 Methylobacter, 278 Methylobacterium, 261 Methylocaldum, 275 Methylomicrobium, 278 Methylomonas, 277 Myxobacteria, 22–23 Neisseria, 605, 622–23 Neisseriaceae, 18 Ochrobactrum, 748–49 Phyllobacterium, 748–49 Prosthecomicrobium, 70 Proteobacteria, 3 Simonsiella, 833–35, 833–37 Sphaerotilus, 773 Spirillum, 703, 707–8 Mottled stripe disease, Herbaspirillum, 143 Mouse habitat. See Rodent habitat Mouth. See Oral cavity Mtases, Neisseria, 626–27 Mucociliary defenses, Bordetella, 658 Mucosal surfaces Alysiella, 833 Comamonas, 726 Eikenella corrodens, 840 Neisseria, 603 Simonsiella, 833 Mule, Burkholderia, 849 Murine typhus, Rickettsia, 503–4 Mussels, Aquaspirillum, 711 Mutagenesis, Zymomonas, 212 Mycobacterium, 320 Mycobacterium flavum, 151 Mycobacterium tuberculosis, 371 Mycoplana, 747 Mycoplana rubra, 257 Myxobacteria, 22–23 morphology of, 22 Myxococcales, 22 Myxococcaceae, 22

Myxococcales, 22 Archangiaceae, 22 Cystobacteraceae, 22 Myxococcaceae, 22 Polyangiaceae, 22 Myxococcus, 22 Myxococcus xanthus, 28 Myxospores, Myxobacteria, 22

N Nalidixic acid resistance, Methylobacterium, 262 Nannocystis, 22 Naphthalene degradation, Comamonas, 729 Nata Acetobacter, 176 Acetobacteraceae, 173 Gluconacetobacter, 181 Native hapten polysaccharide, Brucella, 354, 356, 388–89, 391, 403 Nautilia, 25 Neisseria, 18, 602–32, 720, 828 bacteriophages of, 626 cellular structures of, 608–24 characteristics of, 605–6 disease from, 630–32 gonorrhea, 630–31 meningitis, 631–32 epidemiology of, 628–30 genetics of, 624–28 habitats of, 603 identification of, 604–6 isolation of, 603–4 metabolism of, 607–8 morphology of, 605–6 phylogeny of, 602 physiology of, 607–24 preservation of, 607 proteins of, 608–17 restriction and modification systems of, 626–27 species of, 602 strain typing of, 628–29 transformation of, 625–26 Neisseria animalis, 602–3, 609 Neisseria canis, 602–3, 609 Neisseria cinerea, 602–3, 606–9, 613, 622, 624–25 Neisseria denitrificans, 602, 609 Neisseria dentiae, 602 Neisseria elongata, 602, 605, 608 Neisseria flava, 608–9, 613 Neisseria flavescens, 602–3, 608–9 Neisseria gonorrhoeae, 18, 602–3, 606–30, 632 isolation of, 604 morphology of, 605 Neisseria gonorrhoeae 1291, 620 Neisseria gonorrhoeae FA19, 620 Neisseria lactamica, 602–3, 606–7, 609, 613, 622, 624–25 Neisseria macacae, 602–3 Neisseria meningitidis, 18, 602–3, 606–17, 621–32 isolation of, 604

909

Neisseria mucosa, 602–3, 606, 608, 613, 624–25 Neisseria perflava, 603, 608, 613, 624, 626 Neisseria polysaccharea, 602, 606–7, 609, 624 Neisseria sicca, 602–3, 606, 608–9, 613, 624–25 Neisseria subflava, 602–3, 606, 608, 613, 625 Neisseria weaveri, 602 Neisseriaceae, 18, 737, 828, 837 Alysiella, 18 Chromobacterium, 18 Eikenella, 18 Kingella, 18 Neisseria, 18 Simonsiella, 18 Nematodes, Alcaligenes, 677 Neorickettsia, 13, 460–61, 493, 516–17, 547 disease from, 517 ecology of, 517 genetics of, 516–17 habitats of, 516 physiology of, 517 taxonomy of, 516 Neorickettsia helminthoeca, 13, 461, 516 Neorickettsia, Neorickettsia, 460–61 Neorickettsia risticii, 516–17 Neorickettsia sennetsu, 516 Neptunomonas, 21 Neuroretinitis, Bartonella, 462 Neutrophils, 662–63, 664, 667–68 Brucella, 394 Nickel resistance Achromobacter/Alcaligenes strains, 686 Comamonas, 731 Nickel uptake, Brucella, 363 Nitrate reduction Azospirillum, 127 Brucella, 345, 367 Comamonas, 728 phototrophic alphaproteobacteria, 55 Thiobacillus, 817 Nitrates, manufacture of, 861 Nitric oxide reductase, Achromobacter/Alcaligenes strains, 680–81 Nitrilase Achromobacter/Alcaligenes strains, 687–88 Comamonas, 731 Nitrite oxidation, 861, 862 Nitrite oxidoreductase (NOR), 862, 864–65 Nitrite reductases, Achromobacter/Alcaligenes strains, 680–81 Nitrite reduction, Neisseria, 607 Nitrite-oxidizing bacteria, 861–69 biodegradation by, 868–69 biofilms of, 867 dominant members of, 863–65 enumeration of, 867–68 habitat of, 866–67

910

Index

history of, 861 inhibition of, 868–69 isolation of, 865–66 lithotrophy of, 869 nutritional requirements of, 865–66 resistance of, 868–69 systematics of, 862–65 in wastewater, 868 Nitrobacter, 9, 42, 862–64, 866, 868–69 Nitrobacter alkalicus, 866 Nitrobacter hamburgensis, 862, 864, 867–68 Nitrobacter hamburgensis X14, 864 Nitrobacter sp. strain LL, 864 Nitrobacter vulgaris, 868 Nitrobacter winogradsky, 864–65, 868 Nitrobacter winogradsky serotype agilis, 868 Nitrobacteraceae, 862 Nitrococcus, 862–63, 865 Nitrogen cycle, 861 Nitrogen fixation Alphaproteobacteria, 13–14 Azoarcus, 877, 885–86 Azonexus, 889–90 Azospira, 889–90 Azospirillum, 115, 120, 122, 126–28, 132–34 Azovibrio, 889–90 Beijerinckia, 151, 154, 158–60 Betaproteobacteria, 15 Derxia, 755–56 Gluconacetobacter, 180, 184–85 Herbaspirillum, 141, 143–47 methanotrophs, 283–84 Methylocystis, 280 Methylomonas, 277 Methylosinus, 280 Methylosphaera, 279 phototrophic alphaproteobacteria, 57–58 Phyllobacterium, 749 Xanthobacter, 293, 295–97, 306 Nitrogen metabolism, methanotrophs, 283–84 Nitrogenase, Azoarcus, 885–86 Nitrosococcus, 19, 780, 783, 786– 806 Nitrosococcus halophilus, 783, 786, 803–4 Nitrosococcus mobile, 720 Nitrosococcus mobilis, 783–84, 786, 790, 801, 804–5 Nitrosococcus nitrosus, 786 Nitrosococcus oceani, 783, 786, 803–4 Nitrosococcus oceanus, 862 Nitrosolobus, 720, 780, 783, 785– 806 Nitrosolobus multiformis, 786, 806 Nitrosomonas, 17, 780, 782, 786–806, 863–64, 867, 869 Nitrosomonas aestuarii, 785, 802 Nitrosomonas communis, 784, 790, 796, 801–2, 806

Nitrosomonas cryotolerans, 785, 802, 804–5 Nitrosomonas europaea, 720, 778, 783–84, 790, 795, 801, 804, 864, 867 Nitrosomonas eutropha, 783–84, 790, 801, 804–5 Nitrosomonas halophila, 783–84, 801, 804–5 Nitrosomonas marina, 784, 790, 793, 796, 802–4 Nitrosomonas nitrosa, 784, 786, 801–2, 805 Nitrosomonas oligotropha, 784, 790, 793, 796, 802, 804–6 Nitrosomonas ureae, 784, 802 Nitrosospira, 17, 720, 780, 782, 785–806, 867 Nitrosospira briensis, 785 Nitrosovibrio, 720, 780, 782, 785–806 Nitrospina, 862–65, 867 Nitrospira, 862–63, 865, 868 Nitrospira marina, 863 Nitrospira moscoviensis, 862–63, 865 Nitrous Oxide (N2O) reductase, Achromobacter/Alcaligenes strains, 680–81 Normal human serum Burkholderia resistance to, 855– 56 Neisseria resistance to, 618–19 Novosphingobium, 14 Nutrient uptake, Brucella, 362–63, 408 Nutritional requirements, Brucella, 338–41

O O-antigen Bordetella, 661 Burkholderia, 855 Oceanospirillum, 21, 701, 710, 719 Ocherous masses, Leptothrix, 764–65 Ochrobactrum, 315–17, 321, 324, 330, 365, 370, 747–49 applications of, 749 differentiating features of, 748 habitat of, 748 identification of, 748–49 isolation of, 748 motility of, 748–49 physiology of, 749 preservation of, 748 Ochrobactrum anthropi, 747–49 Ochrobactrum intermedium, 315, 320, 329, 352, 361 Octadecabacter, 14, 565 Oil environment, Xanthobacter, 295, 298 Oligella, 675, 691 Oligella ureolytica, 691 Oligella urethralis, 691 Oligosaccharides Brucella, 352, 360 Neisseria, 617–18

Opine production, Agrobacterium, 94–96, 102, 104 Oral cavity Alysiella, 832–34, 837 Eikenella corrodens, 840 Neisseriaceae, 18 Simonsiella, 829, 832–34, 837 Orangutan Alysiella, 832–33 Simonsiella, 832–33 Organic acid utilization Acidomonas, 179 Azospirillum, 115, 121, 123 Gluconacetobacter, 181 Herbaspirillum, 145–46 Methylobacterium, 257 Paracoccus, 240–43 phototrophic alphaproteobacteria, 55, 56 Orientia, 13, 493, 506–7 disease from, 507 epidemiology of, 507 genetics of, 506–7 habitats of, 506 identification of, 506 isolation of, 506 phylogeny of, 506 physiology of, 506 preservation of, 506 treatment of, 507 Orientia tsutsugamushi, 459–60, 505–7 Ornibactin, Burkholderia, 853–54 Oroya fever, Bartonella, 462, 479, 482–84 Orthanilic acid degradation, Achromobacter/Alcaligenes strains, 684 Osmotolerance, Azospirillum, 131 Outer membrane, 610 Anaplasma, 509 Bartonella, 471 Brucella, 357–61, 402–3, 408 dimorphic prosthecate bacteria, 74, 82 Ehrlichia, 512 Neisseria, 609, 613, 616–17 Rickettsia, 498 Outer-membrane proteinmacromolecular complex, Neisseria, 617 Ovine habitat Brucella, 335, 372–82, 429 Campylobacteraceae, 24–25 Coxiella, 530, 536 Simonsiella, 829 Succinivibrionaceae, 19–20 Oxalobacter, 16–17, 21 Oxalobacter formigenes, 17 Oxygen gradients, Thiobacillus, 812 Oxygen requirements aerobic phototrophic bacteria, 575–78 Derxia, 755 Neisseria, 607 nitrite-oxidizing bacteria, 866–67 Spirillum, 707 Oxygen tolerance, Azoarcus, 885

Index

P Palm wine Acetobacter, 176 Acetobacteraceae, 169 Gluconobacter, 186 Zymomonas, 201–2 Pantothenic acid requirement, Acidomonas, 179 Paracoccus, 14, 232–46, 754 biological potential of, 246 characteristics of, 233–35 chromosomes and plasmids of, 245–46 ecology and isolation of, 237–40 metabolism of, 244–45 phylogenetic relationships in, 236–37 phylogenetic tree of, 236–37 species of, 235–37 Paracoccus alcaliphilus, 234, 240 Paracoccus alkenifer, 236, 240 Paracoccus aminophilus, 235, 240–41 Paracoccus aminovorans, 235, 241 Paracoccus carotinifaciens, 236, 238, 241 Paracoccus denitrificans, 14, 42, 232–46, 351, 680 Paracoccus halodenitrificans, 232 Paracoccus kocurii, 235, 238, 241 Paracoccus kondratievae, 241–42 Paracoccus marcusii, 236, 238, 242 Paracoccus methylutens, 235–36, 242 Paracoccus pantotrophus, 232, 234, 236, 238, 242, 244–46, 812, 816–17, 821 Paracoccus solventivorans, 236, 242–43 Paracoccus thiocyanatus, 234, 238, 243 Paracoccus versutus, 232, 234, 236, 238, 243–46, 821 Paracraurococcus, 562, 566 Paracraurococcus ruber, 163, 562, 565 Parasitism bdellovibrio group, 23 Brucella, 324, 329, 332 Pasteurella, 19, 841 Pasteurella multocida, 28 Pasteurellaceae, 19–20 Actinobacillus, 19 Haemophilus, 19 Mannheimia, 19 Pasteurella, 19 Pathogenicity, 654–61 Acetobacter, 178 Agrobacterium, 99–101 Alysiella, 837 Bartonella, 467, 473–78 Bordetella, 648, 650, 652–54, 662, 667–68 Brucella, 315, 337 Burkholderia, 853–56 Chromobacterium, 739 Comamonas, 731–32 Coxiella, 536–41 Eikenella corrodens, 842–44 Herbaspirillum, 143

Neisseria, 603, 609–16, 618–19, 622, 624 Ochrobactrum, 748 Simonsiella, 837 Pathogenicity island theory, 650 Pathogens Achromobacter, 678–80 Alcaligenes, 678–80 Bordetella, 648–68 Burkholderia, 848–56 Chromobacterium, 739 Eikenella corrodens, 840–45 Neisseria, 602–32 Ochrobactrum, 748 Oligella, 691 Pelistega, 690–91 Taylorella equigenitalis, 689–90 Pattern recognition receptors, 663 Pear habitat, Acetobacteraceae, 172–73 Pectobacterium, 19 Pedomicrobium, 15, 72–86 Pelistega, 16, 675, 690–91 Pelistega europaea, 690–91 Pelobacter group, 23–24 Penicillin resistance Methylobacterium, 262 Xanthobacter, 303 Penicillin sensitivity Seliberia, 588 Xanthobacter, 303 Pentose phosphate pathway Brucella, 363 Coxiella, 534 Gluconobacter, 188 Paracoccus, 245 Zymomonas, 213–14 Peptidoglycan Brucella, 349–50, 359–60 Coxiella, 531 dimorphic prosthecate bacteria, 82 Neisseria, 623–24 Phenylobacterium, 254–55 Periodontal disease, Eikenella corrodens, 843–44 Peritoneal fluid, Achromobacter, 678 Pertactin, 665–67 Bordetella, 656 Pertussis, 649–51, 658, 662–67 acquired immunity to, 664–66 clinical diagnosis of, 652–53 immunity against, 662–63 treatment of, 653 Pertussis toxin, 664–67 B subunit activities of, 659–60 binding of, 659–60 Bordetella, 650–51, 653, 655, 658–60 inactivation of, 659–60 secretion of, 658–59 pH tolerance Acidomonas, 179 aerobic phototrophic bacteria, 565–66 Azospirillum, 125–26 Beijerinckia, 154, 158–59 Brucella, 338 Coxiella, 534, 540

911

dimorphic prosthecate bacteria, 74 Herbaspirillum, 145 Methylobacter, 278 Methylocystis, 280 Methylomicrobium, 279 Methylomonas, 277 Methylosinus, 281 Methylosphaera, 279 Paracoccus, 233–34, 240–43 phototrophic alphaproteobacteria, 45 Xanthobacter, 299–300 Zymomonas, 201–2 Phaeospirillum, 13–14, 41, 43–44 Phaeospirillum fulvum, 46, 56 Phaeospirillum molischianum, 46 Phagocytes, Brucella, 390–98, 402 Phagocytosis of Bordetella, 662–63, 668 Brucella, 406–9 Rickettsia, 496 Phagolysosome, Coxiella, 534, 540–41 Phagosome inhibition, Brucella, 404–5 Phenanthrene degradation, Comamonas, 729 Phenol degradation Achromobacter/Alcaligenes strains, 683 Comamonas, 728, 730 Phenotypic features Comamonas, 727 Leptothrix-Sphaerotilus group, 759–62 Neisseria, 602 Phenylobacterium, 250–56 characteristics of, 252–53 habitats of, 250 identification of, 252–56 isolation and cultivation of, 250–52 Phospholipids, Brucella, 346, 349, 351 Photobacterium, 19 Photoinhibition, nitrite-oxidizing bacteria, 869 Photorhabdus, 19 Photosynthesis, phototrophic betaproteobacteria, 593–99 Phototrophic alphaproteobacteria, 41–59 applications of, 58–59 characteristics of, 51–53 habitats of, 44–45 identification of, 49–54 isolation of, 45–49 media for, 47–49 procedures for, 47 selective enrichment, 46 phylogenetic tree of, 42–43 phylogeny of, 41–42 physiology of, 54–58 carbon metabolism, 55–56 fermentation, 55 hydrogen metabolism, 58 nitrogen metabolism, 57–58 photosynthesis, 54–55

912

Index

respiration, 55 sulfur metabolism, 56–57 preservation of, 54 taxonomy of, 42–44 Phototrophic betaproteobacteria, 593–99 applications of, 599 habitats and ecology of, 596 identification of, 597–98 isolation of, 596–97 phylogeny of, 593 physiology of, 598–99 preservation of, 598 selective enrichment of, 596–97 taxonomy of, 593–94 Phthalate metabolism, Comamonas, 729–30 Phyllobacterium, 14–15, 321, 747–49 applications of, 749 differentiating features of, 748 identification of, 748–49 isolation of, 748 in leaf nodules, 747 motility of, 748–49 physiology of, 749 preservation of, 748 in rhizosphere, 747 Phyllobacterium myrsinacearum, 749 Phyllobacterium rubiacearum, 749 Phylogenetic tree Acetobacteraceae, 163–65 Alysiella, 831 ammonia-oxidizing bacteria, 787–90 Anaplasmataceae, 508 Ancalomicrobium, 67 Azoarcus, 874 Azospirillum, 116 Bartonella, 468 Bordetella, 649–50 Brucella, 316–17, 321 Comamonas, 724 Herbaspirillum, 142 Leptothrix-Sphaerotilus group, 759 methanotrophs, 267–68 Neisseriaceae, 831 Paracoccus, 236–37 phototrophic alphaproteobacteria, 42–43 phototrophic betaproteobacteria, 594 Prosthecomicrobium, 67 Proteobacteria, 9–10 Rhodobacter, 43 Rhodocyclus/Thauera/Azoarcus group, 874 Rhodopseudomonas, 42 Rhodospirillum, 42 Rickettsia, 495 Rickettsiales, 457–58, 494 Simonsiella, 831 Wolbachia, 548–50 Phylogeny aerobic phototrophic bacteria, 562–65 Agrobacterium, 91 Alphaproteobacteria, 9–15

Alysiella, 828–32 ammonia-oxidizing bacteria, 786–94 Anaplasma, 507–8 Azoarcus, 873–75 Azonexus, 888 Azospira, 888 Azovibrio, 888 Bartonella, 467 Betaproteobacteria, 15–18 Brucella, 315–30 Comamonas, 723 Coxiella, 529–30 Ehrlichia, 512 Gammaproteobacteria, 18–21 Leptothrix-Sphaerotilus group, 758–62 methanotrophs, 266–68 Neisseria, 602 nitrite-oxidizing bacteria, 864 Orientia, 506 phototrophic alphaproteobacteria, 41–42 Proteobacteria, 4–25 Rickettsia, 506 Simonsiella, 828–32 for speciation, 333–34 Wolbachia, 547–50 Xanthobacter, 294 Physiology. See also Biochemistry; Ecophysiology Acetobacter, 176–77 Achromobacter, 679 aerobic phototrophic bacteria, 572–78 Agrobacterium, 94–96 Alcaligenes, 679 Alphaproteobacteria, 9–15 Anaplasma, 509 Asaia, 180 Azoarcus, 879–86 Azonexus, 889–90 Azospira, 889–90 Azospirillum, 126–32 Azovibrio, 889–90 Bartonella, 471–78 Beijerinckia, 154–57, 159–60 Betaproteobacteria, 15–18 Brucella, 349–68 Comamonas, 728–29 Coxiella, 534–35 Derxia, 755–56 Ehrlichia, 512 Gammaproteobacteria, 18–21 Gluconobacter, 186–87 Kerstersia, 679 Kozakia, 190 Leptothrix, 770–72 Leptothrix-Sphaerotilus group, 761 methanotrophs, 281–84 Neisseria, 607–24 Neorickettsia, 517 Ochrobactrum, 749 Orientia, 506 Paracoccus, 233–35 Phenylobacterium, 252–53 phototrophic alphaproteobacteria, 54–58

phototrophic betaproteobacteria, 598–99 Phyllobacterium, 749 Pigmentiphaga, 679 Proteobacteria, 9–25 Rickettsia, 496–98 Sphaerotilus, 774–75 Spirillum, 707 Thiobacillus, 816 Xanthobacter, 304–6 Phytohormone production, Herbaspirillum, 147 Pickles, Acetobacter, 176 Pig habitat. See Porcine habitat Pig manure, Aquaspirillum, 711 Pigeons, Pelistega europaea, 691 Pigment Acidomonas, 179 aerobic phototrophic bacteria, 570–71, 575 Agrobacterium, 92 Asaia, 180 Azospirillum, 124–25 Beijerinckia, 155, 157–59 Methylobacter, 278 Methylobacterium, 257, 261 Methylocystis, 280 Methylomonas, 269, 277 Paracoccus, 240–43 Phenylobacterium, 254 phototrophic alphaproteobacteria, 49 Xanthobacter, 290, 293, 300 Pigmentiphaga, 675, 676, 689 physiology of, 679 Pigmentiphaga kullae, 675, 678, 689 Pili Eikenella corrodens, 842 Neisseria, 605–6, 622–23 Pilin, Neisseria, 622 α-Pinene degradation, Alcaligenes strains, 682, 684 Pink-pigmented facultatively methylotrophic bacteria. See Methylobacterium Piscirickettsia, 20, 462 Piscirickettsia salmonis, 20, 462 Plant disease Agrobacterium, 91–110 Alphaproteobacteria, 14–15 Enterobacteriaceae, 19 Proteobacteria, 11–12 Pseudomonas, 21 Plant habitat Agrobacterium, 91–92 Azoarcus, 873, 875, 886–87 Azospira, 888 Azospirillum, 118–19, 127–28, 132 Azovibrio, 888 Burkholderia, 850 Herbaspirillum, 143, 147 Janthinobacterium, 742 Methylobacterium, 259 Phyllobacterium, 747–48 Rhodocista, 132 Skermanella, 132 Xanthobacter, 295–96

Index Plasmids Achromobacter/Alcaligenes strains, 685–86 Agrobacterium, 91, 96–101 Azoarcus, 886 Azospirillum, 127 Brucella, 321–24, 368, 372 Comamonas, 730–31 conjugative, Neisseria, 625 Coxiella, 530–31, 535 cryptic, Neisseria, 624–25 Gluconacetobacter, 184–85 Gluconobacter, 188 Neisseria, 626 Paracoccus, 245–46 Phenylobacterium, 254 Zymomonas, 212 Pleural fluid Achromobacter, 678 Alcaligenes, 677 Pneumonia, Legionella, 21 Polar environment dimorphic prosthecate bacteria, 76 phototrophic alphaproteobacteria, 45 phototrophic betaproteobacteria, 596 Polar lipids Achromobacter, 676 aerobic phototrophic bacteria, 571 Alcaligenes, 676 Kerstersia, 676 Methylocystis, 280 phototrophic alphaproteobacteria, 50 Pigmentiphaga, 676 Polar membrane, Spirillum, 703 Polaromonas, 723 Poly-3-hydroxyalkanoates, phototrophic alphaproteobacteria, 59 Poly-3-hydroxybutyrate, 682 Alcaligenes faecalis production of, 688 phototrophic alphaproteobacteria, 59 Poly-3-hydroxybutyrate degradation Alcaligenes strains, 682–83 Comamonas, 729 Polyamine pattern, Phenylobacterium, 254–55 Polyangiaceae, 22 Poly-β-hydroxybutyrate, Sphaerotilus, 774 Polycyclic aromatic hydrocarbons degradation Achromobacter/Alcaligenes strains, 683–84 Comamonas, 728 Polymerase chain reaction (PCR) Bordetella, 653 Leptothrix-Sphaerotilus group, 764 nitrite-oxidizing bacteria, 863–64, 868 Polymyxin B resistance/sensitivity Brucella, 360–61

Methylobacterium, 262 Xanthobacter, 303 NH-Polysaccharide. See Native hapten polysaccharide O-Polysaccharide, Brucella, 351–56, 372, 388–89, 402–3, 418, 424 Polysaccharide production Agrobacterium, 92 Azospirillum, 127–28 Polysaccharide utilization Acidomonas, 179 Gluconacetobacter, 181 Poplar habitat, Agrobacterium, 92 Porcine habitat Bordetella, 651 Brucella, 335, 372–82 Chromobacterium, 739 Lawsonia, 23 Porins Brucella, 359 Neisseria, 608–10 Porphyrobacter, 14, 562, 565, 571 Porphyrobacter neustonensis, 566 Porphyrobacter tepidarius, 565 Potomac horse fever, Neorickettsia, 517 Preservation Acetobacteraceae, 175 aerobic phototrophic bacteria, 571 Agrobacterium, 94 Alysiella, 837 ammonia-oxidizing bacteria, 778–79 Anaplasma, 509 Ancalomicrobium, 68–69 Aquaspirillum, 718–19 Azoarcus, 885 Azonexus, 889 Azospira, 889 Azospirillum, 123 Azovibrio, 889 Bartonella, 471 Beijerinckia, 154 Brucella, 348–49 Chromobacterium, 740 Comamonas, 728 Derxia, 753 dimorphic prosthecate bacteria, 85 Ehrlichia, 512 Gluconacetobacter, 181 Herbaspirillum, 144–45 Janthinobacterium, 743 Leptothrix, 769 methanotrophs, 281 Methylobacterium, 263 Neisseria, 607 Ochrobactrum, 748 Orientia, 506 Phenylobacterium, 252 phototrophic alphaproteobacteria, 54 phototrophic betaproteobacteria, 598 Phyllobacterium, 748 Prosthecomicrobium, 68–69 Rickettsia, 496 Simonsiella, 837 Sphaerotilus, 774

913

Spirillum, 706–7 Wolbachia, 552 Xanthobacter, 300 Zymomonas, 202 Prevention gonorrhea, 631 meningococcal disease, 632 Pringsheim proposal, 761–62 Prokaryotic endosymbionts of eukaryotic cell, 463 mitochondria, 463–64 Propionate production, phototrophic alphaproteobacteria, 55 Propionibacterium pentosaceum, 365 Propionivibrio, 888 Prosthecomicrobium, 65–70 characteristics of, 69 cultivation and maintenance of, 68–69 habitats of, 65–66 identification of, 69–70 isolation of, 66–68 morphology of, 65, 70 phylogenetic tree of, 67 species of, 67 taxonomy of, 65 Prosthecomicrobium consociatum, 65, 70 Prosthecomicrobium enhydrum, 65, 70 Prosthecomicrobium hirschii, 65, 70 Prosthecomicrobium litoralum, 70 Prosthecomicrobium mishustinii, 65, 70 Prosthecomicrobium pneumaticum, 65–66, 70 Prosthecomicrobium polyspheroidum, 65, 70 Protaminobacter rubrum, 257 Protease, Achromobacter/ Alcaligenes strains, 688 Proteobacteria α-subclass, 3–29 aerobic phototrophs, 562–80 applications of, 12 β-subclass, 3–29 γ-subclass, 3–29 characteristics of, 8 classification of, 6 δ-subclass, 3–29 ecology of, 9–25 ε-subclass, 3–29 eukaryotic cell-associated, 318–19 genera of, 8 genome analysis, 28 introduction to, 3–29 morphology of, 3, 9–25 motility of, 3 parasites, 25–28 phenotypic groups of, 4 phylogenic tree of, 9–10 phylogeny of, 4–25 physiology of, 9–25 symbionts, 25–28 Pseudoazurins, Achromobacter/ Alcaligenes strains, 680–81

Pseudomonadaceae, 20–21 Azomonas, 21 Azotobacter, 21 Cellvibrio, 21 Pseudomonas, 20–21 Pseudomonas, 6, 18, 20–21, 725, 727, 754, 849, 855 Pseudomonas acidovorans, 725, 728, 751 Pseudomonas acidovorans rRNA branch, 725 Pseudomonas aeruginosa, 18, 20–21, 612, 720, 730–31, 850, 854 Pseudomonas alcaligenes, 725, 727 Pseudomonas azotocolligans, 151 Pseudomonas cepacia, 251, 351 Pseudomonas compransoris, 725 Pseudomonas diminuta, 151 Pseudomonas fluorescens, 21, 685, 747 Pseudomonas fragi, 742 Pseudomonas insolita, 751 Pseudomonas lemoignei, 682 Pseudomonas mesophilica, 257–58 Pseudomonas methanica, 257 Pseudomonas pavonaceae, 861 Pseudomonas pseudoalcaligenes subsp. pseudoalcaligenes, 725, 727 Pseudomonas putida, 729–30 Pseudomonas putida mt-2, 683 Pseudomonas radiora, 257–58 Pseudomonas rhodos, 257 Pseudomonas rubrisubalbicans, 720, 737, 742–43 Pseudomonas saccharophila, 758– 59 Pseudomonas solanacearum, 751 Pseudomonas stutzeri, 680 Pseudomonas syringae, 21 Pseudomonas terrigena, 725 Pseudomonas testosteroni, 725 Psychrobacter, 20 Pulque habitat, Zymomonas, 201 Purple bacteria aerobic phototrophic bacteria, 562–80 Beijerinckia, 151 Proteobacteria, 5, 7, 9 Purple nonsulfur bacteria, 43–45 Alphaproteobacteria, 3 Betaproteobacteria, 15–16 betaproteobacteria, 593–99 Proteobacteria, 5, 7, 9 Purple sulfur bacteria Gammaproteobacteria, 18–19 Proteobacteria, 5 Purple-pigmented bacteria Chromobacterium, 737–41 Janthinobacterium, 737, 741–43 Pus Achromobacter, 678 Comamonas, 726 Pyochelin, Burkholderia, 853 Pyramidon degradation, Phenylobacterium, 250–56 Pyrite oxidation, Thiobacillus, 815 Pyruvate decarboxylase, Zymomonas, 206

Pyruvate dehydrogenase Brucella, 364 Zymomonas, 211–12

Q Q fever Coxiella, 529–30, 536–41 immunity from, 538–39 vaccine for, 538 Quadricoccus, 17 Quinoline degradation, Comamonas, 729 Quinone Methylocystis, 280 Methylosinus, 281 Quorum sensing Agrobacterium, 104 Burkholderia, 854 Vibrionaceae, 19

R Rabbit habitat. See Rodent habitat Ralstonia, 6, 16–18, 731, 849 Ralstonia eutropha, 17, 686 Ralstonia metallidurans, 686 Ralstonia taiwanensis, 17 RDL group. See Rhodocyclus gelatinosus-like group 16S rDNA Acetobacter, 176 Acetobacteraceae, 163–64, 166–67 Acidomonas, 178–79 aerobic phototrophic bacteria, 571 Agrobacterium, 91 Alphaproteobacteria, 9, 13, 15 Asaia, 180 Azoarcus, 873–75, 879–85, 887 Azonexus, 889 Azospira, 889 Azospirillum, 116, 123–26 Azovibrio, 888–89 Betaproteobacteria, 16 Bordetella, 648–49 Brucella, 315 Burkholderia, 849 Comamonas, 723–24 Enterobacteriaceae, 19 Gammaproteobacteria, 18, 20 Gluconacetobacter, 181 Gluconobacter, 186 Herbaspirillum, 141–42 nitrite-oxidizing bacteria, 862, 868 phototrophic alphaproteobacteria, 41, 44, 50 phototrophic betaproteobacteria, 593–94 Proteobacteria, 9 23S rDNA Azospirillum, 116 Gammaproteobacteria, 20 Gluconobacter, 186 Herbaspirillum, 142 Red stripe disease, Herbaspirillum, 143 Refinery oil sludge, Azoarcus, 875

Respiratory chain Brucella, 367 Deltaproteobacteria, 21 Respiratory tract Bordetella, 651, 656–57 Cardiobacteriaceae, 20 Eikenella corrodens, 840 innate immune defenses in, 662–64 Legionella, 21 Neisseriaceae, 18 Ochrobactrum, 748 Rhesus monkeys, Neisseria, 603 Rhizobia, Alphaproteobacteria, 3 Rhizobiaceae, 14–15 Agrobacterium, 91 Rhizobium, 14–15 Rhizobiales, Hyphomicrobiaceae, 294 Rhizobium, 13–15, 134, 151, 315, 321, 329, 350, 467–68, 747 Rhizobium loti, 365 Rhizobium meliloti, 28, 97, 371–72 Rhizobium phaseoli, 91 Rhizobium tropicii, 365 Rhizomonas, 203 Rhizosphere Agrobacterium, 105 Azospirillum, 122–23, 127, 131 Beijerinckia, 161 Burkholderia, 850 Gluconacetobacter, 173, 181, 184 Phyllobacterium, 747–48 Xanthobacter, 295, 297 Rhodobaca bogoriensis, 50 Rhodobacter, 9, 13–14, 41–42, 44, 56, 236 characteristics of, 53 phylogenetic tree of, 43 Rhodobacter azotoformans, 55 Rhodobacter blastica, 44, 56 Rhodobacter capsulatus, 14, 56–58, 565, 578, 596, 680 Rhodobacter sphaeroides, 14, 28, 50, 56–59, 96, 565, 571, 573, 578, 580 Rhodobacter veldkampii, 57 Rhodobium, 15, 41, 44–45, 50 Rhodobium marinum, 44–45, 49–50, 56–57 Rhodobium orientis, 44–45, 55 Rhodoblastus, 44 Rhodoblastus acidophilus, 44–46, 56 Rhodocista, 116 Rhodocista centenaria, 42–44, 50, 126, 132 Rhodocyclales, 873 Rhodocyclus, 16, 17, 593–95, 597–98, 888 characteristics of, 595 Rhodocyclus gelatinosus, 594 Rhodocyclus gelatinosus-like (RDL) group, 594 Rhodocyclus purpureus, 17, 43, 56, 594, 596–98 Rhodocyclus tenuis, 56, 593–98 Rhodoferax, 16, 593, 596–97, 599, 723 characteristics of, 595

Index Rhodoferax antarcticus, 594, 596–99 Rhodoferax fermentans, 593–94, 596–99 Rhodomicrobium, 15, 41, 44, 65 Rhodomicrobium vannielii, 44–46, 50, 56–57 Rhodopila, 13, 41, 43–44 Rhodopila globiformis, 41, 43, 45, 54, 57, 163, 565 Rhodoplanes, 44 Rhodoplanes elegans, 44, 55–56 Rhodoplanes roseus, 44, 55–56 Rhodopseudomonas, 15, 41, 43–44, 65, 151, 864 characteristics of, 52 phylogenetic tree of, 42 Rhodopseudomonas acidophilia, 50 Rhodopseudomonas cryptolactis, 44 Rhodopseudomonas gelatinosa, 594 Rhodopseudomonas julia, 44, 57 Rhodopseudomonas palustris, 42, 44–46, 56–58, 598, 862, 864 Rhodospira, 13–14, 43 Rhodospira trueperi, 44–45, 50 Rhodospirillaceae, 13–14 Phaeospirillum, 13–14 Rhodospira, 13–14 Rhodospirillum, 13–14 Rhodovibrio, 13–14 Rhodospirillum, 9, 13–14, 41, 43, 703 characteristics of, 51 phylogenetic tree of, 42 Rhodospirillum photometricum, 44, 46 Rhodospirillum rubrum, 44, 46, 49–50, 55–59, 128, 306, 571, 573, 720 Rhodospirillum tenue, 594 Rhodothalassium, 41, 43–44 Rhodothalassium salexigens, 41, 45–46, 54, 56 Rhodovibrio, 13–14, 41, 43–44 Rhodovibrio salinarum, 45–46, 54 Rhodovibrio sodomensis, 45–46, 50, 54 Rhodovulum, 14, 42, 44–45, 50, 56 Rhodovulum adriaticum, 45–46, 57 Rhodovulum euryhalinum, 45, 57 Rhodovulum iodosum, 44 Rhodovulum robiginosum, 44 Rhodovulum strictum, 56–57 Rhodovulum sulfidophilum, 45–46, 57 Rice habitat Azoarcus, 875 Azonexus, 888 Azospira, 888 Azospirillum, 122, 126–27 Azovibrio, 888 Herbaspirillum, 143, 147 Methylobacterium, 258 Xanthobacter, 295–97 Rickettsia, 13, 315, 320, 323, 457–60, 493, 495–506 diagnosis of, 504–5 disease from, 456–60, 500–504 ecology of, 500 epidemiology of, 500 genetics of, 498–500

habitats of, 496, 501 identification of, 496 isolation of, 496 morphology of, 457–59 phylogenetic tree of, 495 phylogeny of, 495 physiology of, 496–98 host cell invasion, 496–97 host cell release, 497 reactivation of, 497–98 preservation of, 496 spotted fever group, 495 taxonomy of, 495–96 treatment for, 505–6 typhus group, 495 Rickettsia aeschlimannii, 459, 495 Rickettsia africae, 459, 495, 500, 503, 505 Rickettsia akari, 459, 495, 499–500, 503 Rickettsia amblyommii, 500 Rickettsia australis, 459, 495, 499–500, 505 Rickettsia bellii, 495–96, 500 Rickettsia canadensis, 495–96 Rickettsia conorii, 459, 495, 497–500, 503, 505 Rickettsia felis, 459, 495, 500, 504–5 Rickettsia helvetica, 459, 495, 500, 505 Rickettsia honei, 459, 495, 500, 505 Rickettsia japonica, 459, 495–96, 500, 505 Rickettsia massiliae, 459, 495 Rickettsia mongolotimonae, 459 Rickettsia montanensis, 459, 495, 500 Rickettsia parkeri, 459, 495 Rickettsia peacockii, 500 Rickettsia prowazekii, 28, 459, 464, 495, 497–500, 504–5 Rickettsia rhipicephali, 459, 495, 500 Rickettsia rickettsii, 459, 496–500, 505 Rickettsia sibirica, 459, 495, 500, 505 Rickettsia slovaca, 459, 495, 500, 505 Rickettsia typhi, 459, 495, 497, 500, 504–5 Rickettsiaceae, 13, 457–60, 493 Orientia, 495 Rickettsia, 13, 495 Rickettsiales, 457–64, 493–517 Anaplasmataceae, 493, 547 Bartonellae, 462 Chlamydiae, 463 classification of, 459 Ehrlichia group, 460–61 Holospora group, 462 Neorickettsia group, 461 phylogenetic tree of, 457–58, 494 Rickettsia group, 457–60 Rickettsiaceae, 493 Thiomicrospira, 462–63 Wolbachia group, 461–62 Rickettsialpox, Rickettsia, 503–4 Rickettsieae, Rickettsia, 457–60 Rickettsiella, 21, 462 Rickettsiella grylli, 462–63, 493, 529 Rickettsioses, 500–504

915

Rifampicin sensitivity Brucella, 428–29 Orientia, 507 RNA-RNA hybridization, Herbaspirillum, 141 Rochalimaea, 15, 529. See also Bartonella Rocks, nitrite-oxidizing bacteria, 866, 869 Rocky mountain spotted fever, Rickettsia, 502–3 Rodent habitat Bartonella, 469, 479 Brucella, 335, 381–82, 386, 394, 411–12 Chromobacterium, 739 Comamonas, 726 Coxiella, 529–30, 532, 536, 539 Neisseria, 603 Root habitat. See also Grass roots Azospirillum, 118–23, 126–28, 132–34 Herbaspirillum, 143–44, 147 Rose habitat, Agrobacterium, 92 Roseateles, 16, 562, 758 Roseateles depolymerans, 565 Roseivivax, 562, 565 Roseivivax halodurans, 566 Roseivivax halotolerans, 566 Roseobacter, 14, 562, 565, 571–72 Roseobacter algicola, 569 Roseobacter denitrificans, 571, 573, 575–76 Roseobacter gallaeciensis, 569 Roseococcus, 562, 570–72, 577 Roseococcus thiosulfatophilus, 163, 565, 569–73, 575–76, 578, 580 Roseospira, 43–44 Roseospira mediosalina, 45, 50, 57 Roseospirillum, 43 Roseospirillum parvum, 44–45, 50 Roseovarius, 14, 562, 565 Roseovarius tolerans, 566 5S rRNA Acetobacteraceae, 163 Acidomonas, 178 Rickettsiales, 493 Wolbachia, 547 16S rRNA Acetobacter, 176 Acetobacteraceae, 163 aerobic phototrophic bacteria, 565 Alysiella, 828–32 ammonia-oxidizing bacteria, 785–87, 789–806 Ancalomicrobium, 65 Aquaspirillum, 719 Asaia, 180 Azoarcus, 873 Azospirillum, 117, 124 Bartonella, 467–68 Burkholderia, 849 Coxiella, 529 Ehrlichia, 460 Gluconacetobacter, 181 Herbaspirillum, 141–42, 146 history of, 5–6 Leptothrix-Sphaerotilus group, 758–59, 763

916

Index

methanotrophs, 266–68, 271–72 nitrite-oxidizing bacteria, 864 Paracoccus, 233, 236 Phenylobacterium, 254–55 Prosthecomicrobium, 65 Proteobacteria, 5–9 Rickettsia, 495–96 Rickettsiales, 457, 493–94 Simonsiella, 828–32 Wolbachia, 547 23S rRNA ammonia-oxidizing bacteria, 789 Proteobacteria, 6–7 Rickettsiales, 493 Wolbachia, 547 rRNA superfamily III, 751 Aquaspirillum species, 719–20 Chromobacterium, 737 Comamonas, 725 Janthinobacterium, 737 rRNA superfamily IV, Aquaspirillum species, 720–21 Rubrimonas, 14, 562, 565 Rubrimonas cliftonensis, 566 Rubrivivax, 16, 44, 593, 596–99, 758–59 characteristics of, 595 Rubrivivax gelatinosus, 46, 56, 58, 593–94, 596–99, 675 Rubrivivax group, characteristics of, 760 Ruegeria, 14 Ruegeria algicola, 569 Ruminant habitat Anaplasmataceae, 493 Brucella, 335 Ruminobacter, 19–20

S Sagitulla, 14 Sake Acetobacter, 176 Acetobacteraceae, 169 Salicylic acid, Burkholderia, 853 Salmonella, 19, 650 Salpingitis, Neisseria, 631 Sandaracinobacter, 562, 565, 572, 577 Sandaracinobacter sibiricus, 571–73, 575–77 Scrub typhus Orientia, 507 Rickettsia, 459–60 Seal habitat, Brucella, 335 Sediment environment Azoarcus, 875–76 dimorphic prosthecate bacteria, 76 methanotrophs, 270–72 Methylobacterium, 258 phototrophic alphaproteobacteria, 44–45 Thiobacillus, 813 Xanthobacter, 295 Selenomonas, 703 Seliberia, 585–88 habitats of, 585

identification of, 588 isolation of, 585–88 enrichment, 586 from soil, 586 from water, 586–88 morphology of, 585, 587 Seliberia carboxydohydrogena, 585 Seliberia stellata, 585, 587–88 Sennetsu neorickettsiosis, Neorickettsia, 517 Serogroups, Neisseria meningitidis, 628–29 Serology, Phenylobacterium, 255 Serovar typing, Neisseria, 628–29 Sewage environment Acidomonas, 178 Alcaligenes, 684 Alcaligenes strains, 685 ammonia-oxidizing bacteria, 783, 785, 805 Aquaspirillum, 711 bdellovibrio group, 23 dimorphic prosthecate bacteria, 76, 86 Leptothrix, 765 methanotrophs, 270–72 nitrite-oxidizing bacteria, 866 phototrophic alphaproteobacteria, 45, 58–59 phototrophic betaproteobacteria, 596 Seliberia, 585 Xanthobacter, 295 Sewage treatment. See Wastewater treatment Sheath formation Leptothrix, 766–69 Leptothrix-Sphaerotilus group, 759–63 Sphaerotilus, 773–74 structure and composition of Leptothrix, 771–72 Sphaerotilus, 774–75 Sheep habitat. See Ovine habitat Shewanella, 19 Shigella, 19, 650 Sialic acid, Bordetella binding of, 659 Siderophores, 664 Achromobacter/Alcaligenes strains, 688 Azospirillum, 131–32 Burkholderia, 853–54 Neisseria, 613–14 Sieved-soil plate method, 752 Silicibacter, 14 Simonsiella, 18, 828–38 differential traits of, 832 ecology of, 837–38 evolution of, 829–30 habitat of, 832–34 identification of, 835–37 isolation of, 834–35 morphology of, 828, 830, 833–36 in oral cavities of warm-blooded vertebrates, 829 phylogeny of, 828–32 preservation of, 837 taxonomy of, 832

Simonsiella crassa, 829–32, 836–37 Simonsiella muelleri, 829, 832, 836–37 Simonsiella sp. ATCC 27381, 830, 836 Simonsiella steedae, 829–30, 832–33, 836–37 Simonsiella strain ATCC 29437, 836 Sinorhizobium, 14–15, 321, 324, 350 Sinorhizobium fredii, 365 Sinorhizobium meliloti, 91, 96, 324, 365 Skermanella, 116 morphology of, 126 Skermanella parooensis, 126, 132 Slime production Leptothrix-Sphaerotilus group, 759 Sphaerotilus, 775 Xanthobacter, 309 Sludge flocs, nitrite-oxidizing bacteria, 864 Smithella, 24 Soda lakes nitrite-oxidizing bacteria, 866, 869 Thiobacillus, 815 Soft drinks Acetobacteraceae, 173 Gluconobacter, 186 Soil environment Achromobacter, 678 aerobic phototrophic bacteria, 562, 566 Agrobacterium, 91–92, 105 Alcaligenes, 677 Alcaligenes strains, 682 ammonia-oxidizing bacteria, 783–86, 802, 805–6 Ancalomicrobium, 66 Aquaspirillum, 711 Azoarcus, 873, 875–76, 886–87 Azospirillum, 118–19, 122, 131 bdellovibrio group, 23 Beijerinckia, 151–52, 158–61 Burkholderia, 848, 850 Chromobacterium, 738 Comamonas, 726 Derxia, 751–52 dimorphic prosthecate bacteria, 76–77, 83 Janthinobacterium, 741–42 methanotrophs, 270–72 Methylobacterium, 258–59 Myxobacteria, 22 nitrite-oxidizing bacteria, 863, 864 Phenylobacterium, 250 phototrophic alphaproteobacteria, 44 Prosthecomicrobium, 66 Seliberia, 585–86 Thiobacillus, 813, 815 Xanthobacter, 295 L-Sorbose production, Gluconobacter, 188–89 Species classification of, 333–37 generation of, 332–33 Sphaerotilus, 16, 759, 771–75 enrichment of, 772–73

Index habitat of, 772 identification of, 773–74 isolation of, 772–73 morphology of, 773–74 physiology of, 774–75 preservation of, 774 sheath structure and composition of, 774–75 Sphaerotilus discophorus, 761 Sphaerotilus natans, 758–59, 761–62, 768, 771–75 Sphingobium, 14 Sphingomonadaceae, 14 Erythrobacter, 14 Erythromicrobium, 14 Novosphingobium, 14 Porphyrobacter, 14 Sphingobium, 14 Sphingomonas, 14 Sphingopyxis, 14 Sphingomonas, 9, 14, 18, 203, 565 Sphingopyxis, 14 Spinal fluid Achromobacter, 678 Ochrobactrum, 748 Spiral form, 720–21 Spirillum, 701–8, 710, 719 applications of, 707–8 cultivation of, 703–6 differential characteristics of, 704–5 habitat of, 701 identification of, 703 isolation of, 701–3 media for, 702–3 morphology of, 703 motility of, 703, 707–8 physiology of, 707 preservation of, 706–7 selective enrichment of, 701–2 Spirillum minus, 701 Spirillum pleomorphum, 701, 711, 717 Spirillum pulli, 701 Spirillum volutans, 17, 701–8, 710, 720 Spotted fever, Rickettsia, 459, 495–96, 501–2 Sputum Alcaligenes, 677 Aquaspirillum, 711 Burkholderia, 850 Stainer-Scholte broth, 652 Stainless steel corrosion, LeptothrixSphaerotilus group, 763 Staleya, 14 Staphylococcus, 688, 850 Staphylococcus aureus, 667, 688 Starkeya novella, 812, 821 Stenotrophomonas, 21, 727 Stenotrophomonas maltophilia, 21, 850 Steroid degradation, Comamonas, 729–31 Stigmatella, 22 Stone, nitrite-oxidizing bacteria, 866, 869 Strain typing, Neisseria, 628–29 Stratified lakes, Thiobacillus, 812

Streptococcus, 688 Streptococcus anginous, 843 Streptococcus constellatus, 843 Streptococcus intermedius, 843 Streptomycin sensitivity Brucella, 428 Methylobacterium, 262 Succinate production, phototrophic alphaproteobacteria, 55 Succinate utilization, Bartonella, 471 Succinivibrio, 19–20 Succinivibrionaceae, 19–20 Anaerobiospirillum, 19–20 Ruminobacter, 19–20 Succinivibrio, 19–20 Succinomonas, 19–20 Succinoglycan synthesis, Agrobacterium, 97 Succinomonas, 19–20 Sugar tolerance, Zymomonas, 208–9 Sugarcane habitat Acetobacteraceae, 173 Azospirillum, 132 Gluconacetobacter, 180, 181, 184 Herbaspirillum, 143 Zymomonas, 201 Sulfate- and sulfur-reducing bacteria, 23–24 Sulfate reduction, phototrophic betaproteobacteria, 598 Sulfide gradients, Thiobacillus, 812 Sulfide oxidation, Thiobacillus, 820 Sulfitobacter, 14 Sulfolobus, 815 Sulfonic acid degradation, Alcaligenes, 684 Sulfur compound utilization Campylobacteraceae, 25 Paracoccus, 235, 237–39, 244–45 phototrophic alphaproteobacteria, 56–57 Sulfur cycle, 813, 815 Sulfur oxidation, Thiobacillus, 816, 820 Sulfur oxidizers. See Thiobacillus Sulfurospirillum, 25 Sulfurospirillum arsenophilum, 25 Sulfurospirillum barnesii, 25 Superoxide dismutase production Coxiella, 541 dimorphic prosthecate bacteria, 74, 77 Neisseria, 607 Suttonella, 20 Suttonella indologenes, 20 Swamp environment, methanotrophs, 271–72 Swarming Azospirillum, 123 dimorphic prosthecate bacteria, 73, 81 Myxobacteria, 23 Swine habitat. See Porcine habitat Swine waste, Dechlorosoma suillum, 888 Synthrophobacter, 23–24 Syntrophus, 24

917

T T DNA, Agrobacterium, 102–6, 110 Tanning process, Acetobacteraceae, 174 Tartrate utilization, Agrobacterium, 106 Taurine degradation, Achromobacter/Alcaligenes strains, 684 Taxonomy Acetobacter, 175–76 Acetobacteraceae, 164–66 Acidomonas, 178 aerobic phototrophic bacteria, 562–65 Agrobacterium, 91–92 Alysiella, 832 Anaplasmataceae, 508 Ancalomicrobium, 65 Aquaspirillum, 719–21 Asaia, 180 Azoarcus, 875 Azonexus, 888 Azospira, 888 Azospirillum, 115–16 Azovibrio, 888 Bartonella, 467–69 Brucella, 330–37 Burkholderia, 849–50 Comamonas, 723–26 Ehrlichia, 512 Gluconacetobacter, 180–81 Gluconobacter, 185–86 Herbaspirillum, 141–43 Kozakia, 189–90 Leptothrix-Sphaerotilus group, 758–62 methanotrophs, 266, 268–70 Methylobacterium, 257–58 Neorickettsia, 516 Paracoccus, 233–35 phototrophic alphaproteobacteria, 42–44 phototrophic betaproteobacteria, 593–94 Prosthecomicrobium, 65 Rickettsia, 495–96 Simonsiella, 832 for speciation, 334 Wolbachia, 550–51 Xanthobacter, 294–95 Taylorella, 16, 675, 689–90 Taylorella asinigenitalis, 689–90 Taylorella equigenitalis, 689–90 Tellurite reduction, aerobic phototrophic bacteria, 578–80 Temperature tolerance Acidomonas, 179 aerobic phototrophic bacteria, 565–66 Agrobacterium, 92 Azospirillum, 125–26 Beijerinckia, 154, 156, 158–59 Brucella, 338 dimorphic prosthecate bacteria, 74 Herbaspirillum, 145 Methylobacter, 278

918

Index

Methylobacterium, 259–60, 262 Methylococcus, 275 Methylocystis, 280 Methylomicrobium, 279 Methylomonas, 277 Methylosinus, 281 Methylosphaera, 279 Paracoccus, 233–34, 240–43 Xanthobacter, 299–300 Zymomonas, 201–2 Tequila Acetobacter, 176 Acetobacteraceae, 169 Terephthalate degradation, Comamonas, 729, 730 Termobacterium mobile. See Zymomonas mobilis Terpene degradation, Comamonas, 729 Testosterone degradation, Comamonas, 729 Tetracycline resistance/sensitivity Brucella, 428–29 Coxiella, 537 dimorphic prosthecate bacteria, 74 Methylobacterium, 262 Neisseria, 625, 626 Tetrathionate, 813 Th1 pathway, 665–66 Th2 pathway, 665–66 Thauera, 17, 873–75, 877, 888 Thauera aromatica, 886 Thauera selenatis, 17 Thermithiobacillus, 18 Thermodesulforhabdus, 24 Thermothiobacillus tepidarius, 816 Thioalkalimicrobium, 20 Thioalkalivibrio, 19 Thiobacilli. See Thiobacillus Thiobacillus, 17–18, 812–24 bacteria of, 821–24 basic characteristics of, 814 betaproteobacteria bacteria from, 812–14, 821–22 enrichment, isolation, and cultivation of, 816–19 gammaproteobacteria bacteria from, 812–14, 821–24 habitat of, 812–16 identification of, 820–21 physiology of, 816 reorganization of, 812–14, 820 Thiobacillus A2, 821 Thiobacillus acidophilus, 565, 812, 821 Thiobacillus aquaesulis, 821 Thiobacillus dentrificans, 680, 816, 819, 821 Thiobacillus ferrooxidans, 823 Thiobacillus neapolitanus, 821 Thiobacillus novellus, 812, 821 Thiobacillus organoparus, 821 Thiobacillus prosperus, 824 Thiobacillus rapidicrescens, 821 Thiobacillus rubellus, 822 Thiobacillus strain Q, 817, 822 Thiobacillus strains O and W, 816, 819

Thiobacillus thioparus, 17–18, 701, 817, 819, 821 Thiobacillus versutus, 812, 821 Thiodendron, 72–86 Thiomargarita, 20 Thiomicrospira, 462–63 Coxiella, 462 Piscirickettsia, 462 Rickettsiella, 462 Thiomicrospira, 20, 812, 815 Thiomicrospira denitrificans, 25 Thiomonas, 812, 821–22 Thiomonas cuprina, 821–22 Thiomonas intermedia, 822 Thiomonas perometabolis, 822 Thiomonas plumbophilus, 822 Thiomonas thermosulfata, 822 Thioploca, 20 Thiosphaera pantotropha, 821. See also Paracoccus pantotrophus Thiosphera pantotropha, 680, 812 Thiosulfate utilization aerobic phototrophic bacteria, 578 Paracoccus, 233–35, 238, 244 Thiobacillus, 817–18, 820 Thiothrix, 20 Thiovulum, 25, 815 Ti plasmid, Agrobacterium, 96, 100–107, 110 Toluene degradation, Azoarcus, 886 4-Toluene sulfonate degradation, Comamonas, 729, 730 Toluidine degradation, Achromobacter/Alcaligenes strains, 683 Toxin. See also Adenylate cyclase toxin; Dermonecrotic toxin; Pertussis toxin; Tracheal cytotoxin Bordetella adhesin as, 657 Bordetella pertussis, 658–61 Burkholderia, 855 Trachea, Bordetella, 656–57, 667 Tracheal colonization factor, Bordetella, 656–57 Tracheal cytotoxin, 658 Transferrin, 613–14, 663–64 Neisseria binding of, 614 Transmission of Alysiella, 837–38 Burkholderia, 851–53 of Simonsiella, 837–38 Trench fever, Bartonella, 462, 479, 482–84 Tricarboxylic acid cycle Brucella, 363–64 Coxiella, 534 Epsilonproteobacteria, 24 Gluconacetobacter, 183 Gluconobacter, 186–87 Rickettsia, 498 Trichloroethene degradation, Alcaligenes strains, 682 Trophoblasts, Brucella, 397–98 Tumors Agrobacterium, 91–92, 102–4 Phyllobacterium, 749 Turbot gills, Janthinobacterium, 742 Turkey coryza, 651

Type III secretion system, Bordetella, 667–68 Type IV secretion system, Rickettsia, 499–500 Typhus, Rickettsia, 459, 495, 501–4

U Ubiquinones Acetobacter, 175–76 Acetobacteraceae, 163, 166 Acidomonas, 179 Alphaproteobacteria, 7–8, 13 Asaia, 180 Betaproteobacteria, 7–8 Brucella, 367 Gammaproteobacteria, 7–8 Gluconacetobacter, 180–81, 183 Gluconobacter, 186–87 Kozakia, 190 Phenylobacterium, 256 Zymomonas, 209–11 Ultramicrobacterium, 141 Ultrastructure aerobic phototrophic bacteria, 571 Brucella, 349–50 Ehrlichia, 460 Undulant fever, Brucella, 372 Urea cycle, Brucella, 366 Urease activity Brucella, 345, 366, 404 Methylobacter, 278 Methylobacterium, 262 Urethritis, Neisseria, 630 Urine Achromobacter, 678 Alcaligenes, 677 Comamonas, 726 Ochrobactrum, 748 Oligella, 691 Urosepsis, Oligella, 691

V Vaccination Anaplasma, 511–12 Brucella, 375–80 Coxiella, 538 Vaccines Neisseria gonorrhoeae, 609, 612–13, 615–16, 631 Neisseria meningitidis, 609, 612–13, 615–16, 632 pertussis, 651–53, 657–60, 662–66 resistance to, 666–67 Variovorax, 16, 723 Variovorax paradoxus, 16, 593 Verruga peruana, Bartonella, 462 Vibrio, 19, 703, 710, 740 Vibrio alcaligenes, 723 Vibrio cholerae, 19 Vibrio cyclosites, 723, 725 Vibrio extorquens, 257 Vibrio neocistes, 723, 725 Vibrio percolans, 723 Vibrio serpens, 710 Vibrio terrigenus, 725

Index Vibrionaceae, 19–20 Photobacterium, 19 Shewanella, 19 Vibrio, 19 Vinegar Acetobacter, 176, 178 Acetobacteraceae, 170–72 Gluconacetobacter, 185 history of, 170–71 production of, 171–72 Violacein, 737, 739, 743 biosynthesis of, 740–41 structure of, 738 Virulence factors Bordetella, 653–54, 656–58 of Bordetella bronchiseptica cluster, 667–68 Burkholderia, 853 Virulence mechanisms Agrobacterium, 99–101 Brucella, 401–11 Coxiella, 538–41 Virulence structures Bartonella, 477 Brucella, 324–29 Vitamin C production, Gluconobacter, 188–89 Vitronectin, 611 Vorticella microstoma, 741

W Wastewater treatment ammonia-oxidizing bacteria, 805 dimorphic prosthecate bacteria, 76, 85–86 Methylobacterium, 263 nitrite-oxidizing bacteria, 863–66, 868–69 Paracoccus, 246 phototrophic alphaproteobacteria, 58–59 phototrophic betaproteobacteria, 599 Spirillum, 707–8 Thiobacillus, 816 Water habitats. See also Freshwater environment; Marine environment Achromobacter, 678 Achromobacter/Alcaligenes strains, 686–87 Alcaligenes, 677 Azoarcus, 876 Burkholderia, 850 Chromobacterium, 738 Comamonas, 726 Janthinobacterium, 741–42 nitrite-oxidizing bacteria, 865 Whale habitat, Brucella, 335 Wheat habitat, Azospirillum, 119–20, 123, 127 Whole cell hybridization, Leptothrix-Sphaerotilus group, 763–64

Whooping cough. See also Pertussis molecular basis of, 662–67 Wine Acetobacter, 176 Acetobacteraceae, 168–69 Gluconobacter, 186 Wolbachia, 13, 315, 461, 493, 515–16, 547–56 applications of, 516, 555–56 for filariasis control, 556 host population suppression, 555–56 modulation of insecttransmitted disease, 556 disease from, 516, 555 ecology of, 515–16, 553–55 cytoplasmic incompatibility, 553 feminization, 554 male killing, 554–55 thelytokous parthenogenesis, 554 genome of, 552 habitats of, 515 identification of, 551–52 isolation of, 551 morphology of, 515, 551 phylogenetic tree of, 548–50 phylogeny of, 547–50 preservation of, 552 taxonomy of, 550–51 Wolbachia group other bacteria, 461–62 Wolbachiae, 461 Wolbachia melophagi, 461–62, 547 Wolbachia persica, 461, 493, 547 Wolbachia pipientis, 461, 463, 494, 515–16, 547–56 Wolbachiae, Wolbachia, 461 Wolinella, 25 Wolinella succinogenes, 25 Wounds Alcaligenes, 677 Aquaspirillum, 711 Ochrobactrum, 748

X Xanthobacter, 290–310, 754 applications of, 307–9 characteristics of, 291 cultivation of, 299–300 cultures of, 309–10 habitats of, 295–97 identification of, 300–306 biochemical properties, 301–3 physiological properties, 304–6 isolation of, 297–99 phylogeny and taxonomy, 294–95 Xanthobacter agilis, 290, 298–99, 302–3 Xanthobacter aminoxidans, 290, 294, 299–301 Xanthobacter autotrophicus, 290–310

919

Xanthobacter flavus, 290–310 Xanthobacter methylooxidans, 290, 303–4 Xanthobacter polyaromaticivorans, 290, 293, 298, 300, 304 Xanthobacter tagetidis, 290, 293, 300–301, 308–9 Xanthobacter viscosus, 290, 294, 299–301 Xanthomonas, 18, 21, 689, 747 Xanthomonas autotrophicus, 151 Xenobiotic compound degradation, Achromobacter/Alcaligenes strains, 684–86 Xenorhabdus, 19 Xylella, 21 Xylella fastidiosa, 21 Xylophilus, 16, 723 Xylophilus ampelinus, 16

Y Yersinia, 19, 28

Z Zavarzinia compransoris, 725 Zeaxanthin dirhamnoside, Xanthobacter, 303 Zinc metalloprotease, Burkholderia, 854 Zoogloea ramigera, 17 Zoonosis Brucella, 372, 374, 380–81 Coxiella, 530, 536 Ehrlichia, 460, 514 Rickettsia, 504 Zymobacter, 20 Zymomonas, 14, 151, 169, 201–14 characteristics of, 202–3 ethanol fermentation, 207–9 genetics and metabolic engineering, 212–14 growth and conservation of, 202 habitats of, 201 identification of, 202–3 isolation of, 201–2 metabolism of, 203–7 aerobic, 209–11 alcohol dehydrogenase isoenzymes, 206–7 carbohydrate transport, 203–4 glycolytic flux and regulation of, 204–6 pyruvate decarboxylase, 206 sugar and ethanol tolerance, 208–9 Zymomonas mobilis, 174, 201–14 Zymomonas mobilis subsp. mobilis, 202 Zymomonas mobilis subsp. pomaceae, 202