The Prokaryotes: Volume 2: Ecophysiology and Biochemistry [vol 2, 3rd ed.] 9780387254920, 0-387-25492-7

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The Prokaryotes Third Edition

The Prokaryotes A Handbook on the Biology of Bacteria Third Edition

Volume 2: Ecophysiology and Biochemistry 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 2 ISBN-10: 0-387-25492-7 ISBN-13: 978-0387-25492-0 e-ISBN: 0-387-30742-7 Print + e-ISBN: 0-387-33491-2 DOI: 10.1007/0-387-30742-7 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.

Essays in Prokaryotic Biology

1.1

How We Do, Don’t and Should Look at Bacteria and Bacteriology carl r. woese

3

Databases

24

1.3

Defining Taxonomic Ranks

29

1.4

Prokaryote Characterization and Identification hans g. trüper and karl-heinz schleifer

58

1.5

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

1.2

wolfgang ludwig, karl-heinz schleifer and erko stackebrandt erko stackebrandt

jörg overmann

80

1.6

Prokaryotes and Their Habitats hans g. schlegel and holger w. jannasch

137

1.7

Morphological and Physiological Diversity stephen h. zinder and martin dworkin

185

1.8

Cell-Cell Interactions

221

1.9

Prokaryotic Genomics b. w. wren

246

1.10

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

261

dale kaiser

x

Contents

1.11

Origin of Life: RNA World versus Autocatalytic Anabolism

275

1.12

Biotechnology and Applied Microbiology

284

1.13

The Structure and Function of Microbial Communities david a. stahl, meredith hullar and seana davidson

299

2.

Symbiotic Associations

günter wächtershäuser eugene rosenberg

Cyanobacterial-Plant Symbioses

331

2.2

Symbiotic Associations Between Ciliates and Prokaryotes hans-dieter görtz

364

2.3

Bacteriocyte-Associated Endosymbionts of Insects paul baumann, nancy a. moran and linda baumann

403

2.4

Symbiotic Associations Between Termites and Prokaryotes

439

2.5

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

475

3.

Biotechnology and Applied Microbiology

3.1

Organic Acid and Solvent Production palmer rogers, jiann-shin chen and mary jo zidwick

511

3.2

Amino Acid Production

756

3.3

Microbial Exopolysaccharides timothy harrah, bruce panilaitis and david kaplan

766

3.4

Bacterial Enzymes wim j. quax

777

3.5

Bacteria in Food and Beverage Production michael p. doyle and jianghong meng

797

3.6

Bacterial Pharmaceutical Products arnold l. demain and giancarlo lancini

812

3.7

Biosurfactants

834

3.8

Bioremediation ronald l. crawford

850

2.1

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

andreas brune

hidehiko kumagai

eugene rosenberg

Contents

xi

3.9

Biodeterioration ji-dong gu and ralph mitchell

864

3.10

Microbial Biofilms

904

dirk de beer and paul stoodley

939

Index

Volume 2 1.

Ecophysiological and Biochemical Aspects

1.1

Planktonic Versus Sessile Life of Prokaryotes kevin c. marshall

1.2

Bacterial Adhesion itzhak ofek, nathan sharon and soman n. abraham

16

1.3

The Phototrophic Way of Life

32

1.4

The Anaerobic Way of Life ruth a. schmitz, rolf daniel, uwe deppenmeier and

86

jörg overmann and ferran garcia-pichel

3

gerhard gottschalk

1.5

Bacterial Behavior

102

1.6

Prokaryotic Life Cycles

140

1.7

Life at High Temperatures

167

1.8

Life at Low Temperatures

210

1.9

Life at High Salt Concentrations

263

1.10

Alkaliphilic Prokaryotes

283

1.11

Syntrophism among Prokaryotes

309

1.12

Quorum Sensing bonnie l. bassler and melissa b. miller

336

Acetogenic Prokaryotes

354

1.13

judith armitage martin dworkin

rainer jaenicke and reinhard sterner siegfried scherer and klaus neuhaus aharon oren

terry ann krulwich

bernhard schink and alfons j.m. stams

harold l. drake, kirsten küsel and carola matthies

xii

Contents

1.14

Virulence Strategies of Plant Pathogenic Bacteria barbara n. kunkel and zhongying chen

421

1.15

The Chemolithotrophic Prokaryotes donovan p. kelly and anne p. wood

441

1.16

Oxidation of Inorganic Nitrogen Compounds as an Energy Source

457

1.17

The H2-Metabolizing Prokaryotes

496

1.18

Hydrocarbon-Oxidizing Bacteria

564

1.19

Cellulose-Decomposing Bacteria and Their Enzyme Systems edward a. bayer, yuval shoham and raphael lamed

578

1.20

Aerobic Methylotrophic Prokaryotes mary e. lidstrom

618

1.21

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

635

1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes ralf rabus, theo a. hansen and friedrich widdel

659

The Denitrifying Prokaryotes

769

Dinitrogen-Fixing Prokaryotes

793

Root and Stem Nodule Bacteria of Legumes michael j. sadowsky and p. h. graham

818

Magnetotactic Bacteria

842

Luminous Bacteria paul v. dunlap and kumiko kita-tsukamoto

863

Bacterial Toxins

893

The Metabolic Pathways of Biodegradation lawrence p. wackett

956

Haloalkaliphilic Sulfur-Oxidizing Bacteria

969

The Colorless Sulfur Bacteria lesley a. robertson and j. gijs kuenen

985

1.23 1.24 1.25 1.26 1.27 1.28 1.29 1.30

1.31

eberhard bock and michael wagner

edward schwartz and bärbel friedrich

eugene rosenberg

derek lovley

james p. shapleigh

esperanza martinez-romero

stefan spring and dennis a. bazylinski

vega masignani, mariagrazia pizza and rino rappuoli

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

Contents

xiii

1.32

Bacterial Stress Response eliora z. ron

1012

1.33

Anaerobic Biodegradation of Hydrocarbons Including Methane friedrich widdel, antje boetius and ralf rabus

1028

1.34

Physiology and Biochemistry of the Methane-Producing Archaea reiner hedderich and william b. whitman

1050 1081

Index

Volume 3 A:

Archaea

1.

The Archaea: A Personal Overview of the Formative Years ralph s. wolfe

3

Thermoproteales

10

3.

Sulfolobales

23

4.

Desulfurococcales

52

5.

The Order Thermococcales

69

The Genus Archaeoglobus

82

2.

6.

harald huber, robert huber and karl o. stetter harald huber and david prangishvili harald huber and karl o. stetter costanzo bertoldo and garabed antranikian patricia hartzell and david w. reed Thermoplasmatales

101

The Order Halobacteriales

113

9.

The Methanogenic Bacteria william b. whitman, timothy l. bowen and david r. boone

165

10.

The Order Methanomicrobiales jean-louis garcia, bernard ollivier and william b. whitman

208

11.

The Order Methanobacteriales adam s. bonin and david r. boone

231

The Order Methanosarcinales

244

7. 8.

12.

harald huber and karl o. stetter aharon oren

melissa m. kendall and david r. boone

xiv

Contents

Methanococcales william b. whitman and christian jeanthon

257

Nanoarchaeota

274

15.

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

281

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

297

The Family Bifidobacteriaceae

322

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

383

Family Propionibacteriaceae: The Genus Propionibacterium

400

1.1.5

The Family Succinivibrionaceae

419

1.1.6

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

430

1.1.7

The Family Streptomycetaceae, Part I: Taxonomy

538

1.1.8

The Family Streptomycetaceae, Part II: Molecular Biology

605

1.1.9

The Genus Actinoplanes and Related Genera

623

1.1.10

The Family Actinosynnemataceae david p. labeda

654

1.1.11

The Families Frankiaceae, Geodermatophilaceae, Acidothermaceae and Sporichthyaceae

13. 14.

1.1.2 1.1.3

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

erko stackebrandt and peter schumann bruno biavati and paola mattarelli

erko stackebrandt and klaus p. schaal 1.1.4

erko stackebrandt, cecil s. cummins and john l. johnson erko stackebrandt and robert b. hespell

peter kämpfer

hildgund schrempf gernot vobis

philippe normand

669

Contents

1.1.12

xv

The Family Thermomonosporaceae: Actinocorallia, Actinomadura, Spirillospora and Thermomonospora

682

1.1.13

The Family Streptosporangiaceae

725

1.1.14

The Family Nocardiopsaceae

754

1.1.15

Corynebacterium—Nonmedical

796

1.1.16

The Genus Corynebacterium—Medical

819

1.1.17

The Families Dietziaceae, Gordoniaceae, Nocardiaceae and Tsukamurellaceae

843

reiner michael kroppenstedt and michael goodfellow michael goodfellow and erika teresa quintana

reiner michael kroppenstedt and lyudmila i. evtushenko wolfgang liebl

alexander von graevenitz and kathryn bernard

michael goodfellow and luis angel maldonado

1.1.18

The Genus Mycobacterium—Nonmedical sybe hartmans, jan a.m. de bont and erko stackebrandt

889

1.1.19

The Genus Mycobacterium—Medical

919

1.1.20

Mycobacterium leprae thomas m. shinnick

934

The Genus Arthrobacter

945

The Genus Micrococcus

961

1.1.23

Renibacterium hans-jürgen busse

972

1.1.24

The Genus Stomatococcus: Rothia mucilaginosa, basonym Stomatococcus mucilaginosus

1.1.21 1.1.22

beatrice saviola and william bishai

dorothy jones and ronald m. keddie miloslav kocur, wesley e. kloos and karl-heinz schleifer

erko stackebrandt

975 983

1.1.25

The Family Cellulomonadaceae erko stackebrandt, peter schumann and helmut prauser

1.1.26

The Family Dermatophilaceae

1002

The Genus Brevibacterium

1013

The Family Microbacteriaceae lyudmila i. evtushenko and mariko takeuchi

1020

1.1.27 1.1.28

erko stackebrandt

matthew d. collins

xvi

1.1.29

Contents

The Genus Nocardioides

jung-hoon yoon and yong-ha park

1099 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 friedrich götz, tammy bannerman and karl-heinz schleifer

1.2.2

The Genus Streptococcus—Oral

jeremy m. hardie and robert a. whiley

5 76

Medically Important Beta-Hemolytic Streptococci

108

1.2.4

Streptococcus pneumoniae

149

1.2.5

The Genus Enterococcus: Taxonomy luc devriese, margo baele and patrick butaye

163

1.2.6

Enterococcus donald j. leblanc

175

The Genus Lactococcus

205

1.2.8

The Genera Pediococcus and Tetragenococcus wilhelm h. holzapfel, charles m. a. p. franz, wolfgang ludwig, werner back and leon m. t. dicks

229

1.2.9

Genera Leuconostoc, Oenococcus and Weissella

267

1.2.10

The Genera Lactobacillus and Carnobacterium walter p. hammes and christian hertel

320

1.2.11

Listeria monocytogenes and the Genus Listeria nadia khelef, marc lecuit, carmen buchrieser, didier cabanes,

404

The Genus Brochothrix

477

The Genus Erysipelothrix

492

1.2.3

1.2.7

p. patrick cleary and qi cheng elaine tuomanen

michael teuber and arnold geis

johanna björkroth and wilhelm h. holzapfel

olivier dussurget and pascale cossart

1.2.12 1.2.13

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

Contents

xvii

The Genus Gemella

511

The Genus Kurthia

519

1.2.16

The Genus Bacillus—Nonmedical ralph a. slepecky and h. ernest hemphill

530

1.2.17

The Genus Bacillus—Insect Pathogens donald p. stahly, robert e. andrews and allan a. yousten

563

1.2.18

The Genus Bacillus—Medical w. edmund farrar and annette c. reboli

609

1.2.19

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

1.2.20

An Introduction to the Family Clostridiaceae jürgen wiegel, ralph tanner and fred a. rainey

654

1.2.21

Neurotoxigenic Clostridia cesare montecucco, ornella rossetto and michel r. popoff

679

1.2.22

The Enterotoxic Clostridia bruce a. mcclane, francisco a. uzai, mariano e. fernandez miyakawa, david lyerly and

698

Clostridium perfringens and Histotoxic Disease

753

The Genera Desulfitobacterium and Desulfosporosinus: Taxonomy

771

The Genus Desulfotomaculum

787

1.2.26

The Anaerobic Gram-Positive Cocci takayuki ezaki, na (michael) li and yoshiaki kawamura

795

1.2.27

The Order Haloanaerobiales

809

The Genus Eubacterium and Related Genera

823

The Genus Mycoplasma and Related Genera (Class Mollicutes)

836

The Phytopathogenic Spiroplasmas jacqueline fletcher, ulrich melcher and astri wayadande

905

1.2.14 1.2.15

matthew d. collins erko stackebrandt, ronald m. keddie and dorothy jones

631

tracy wilkins

1.2.23 1.2.24 1.2.25

1.2.28 1.2.29 1.2.30

julian i. rood

stefan spring and frank rosenzweig friedrich widdel

aharon oren

william g. wade shmuel razin

xviii

Contents

1.3

Firmicutes with Atypical Cell Walls

1.3.1

The Family Heliobacteriaceae michael t. madigan

951

1.3.2

Pectinatus, Megasphaera and Zymophilus

965

The Genus Selenomonas

982

The Genus Sporomusa

991

1.3.3 1.3.4 1.3.5

auli haikara and ilkka helander

robert b. hespell, bruce j. paster and floyd e. dewhirst john a. breznak

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

1002

The Genus Veillonella

1022

1.3.7

Syntrophomonadaceae

1041

2.

Cyanobacteria

2.1

The Cyanobacteria—Isolation, Purification and Identification john b. waterbury

1053

2.2

The Cyanobacteria—Ecology, Physiology and Molecular Genetics

1074

The Genus Prochlorococcus

1099

michael cotta and robert forster 1.3.6

2.3

paul kolenbrander

martin sobierj and david r. boone

yehuda cohen and michael gurevitz 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

xix

65

3.1.2

The Genera Prosthecomicrobium and Ancalomicrobium gary e. oertli, cheryl jenkins, naomi ward, frederick a. rainey, erko stackebrandt and james t. staley

3.1.3

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

72

The Genus Agrobacterium

91

3.1.4

ann g. matthysse

The Genus Azospirillum

115

The Genus Herbaspirillum

141

The Genus Beijerinckia

151

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

163

The Genus Zymomonas

201

The Manganese-Oxidizing Bacteria kenneth h. nealson

222

The Genus Paracoccus

232

The Genus Phenylobacterium

250

3.1.13

Methylobacterium peter n. green

257

3.1.14

The Methanotrophs—The Families Methylococcaceae and Methylocystaceae

266

The Genus Xanthobacter

290

The Genus Brucella

315

Introduction to the Rickettsiales and Other Intracellular Prokaryotes david n. fredricks

457

The Genus Bartonella

467

3.1.5 3.1.6 3.1.7 3.1.8

anton hartmann and jose ivo baldani michael schmid, jose ivo baldani and anton hartmann jan hendrick becking

jean swings 3.1.9 3.1.10 3.1.11 3.1.12

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

donovan p. kelly, frederick a. rainey and ann p. wood jürgen eberspächer and franz lingens

john p. bowman 3.1.15 3.1.16 3.1.17 3.1.18

jürgen wiegel

edgardo moreno and ignacio moriyón

michael f. minnick and burt e. anderson

xx

Contents

3.1.19 3.1.20 3.1.21 3.1.22 3.1.23

The Order Rickettsiales xue-jie yu and david h. walker

493

The Genus Coxiella

529

The Genus Wolbachia

547

Aerobic Phototrophic Proteobacteria

562

The Genus Seliberia

585

robert a. heinzen and james e. samuel markus riegler and scott l. o’neill vladimir v. yurkov

jean m. schmidt and james r. swafford

3.2.

Beta Subclass

3.2.1

The Phototrophic Betaproteobacteria johannes f. imhoff

593

3.2.2

The Neisseria daniel c. stein

602

The Genus Bordetella

648

Achromobacter, Alcaligenes and Related Genera hans-jürgen busse and andreas stolz

675

The Genus Spirillum

701

The Genus Aquaspirillum

710

Comamonas

723

The Genera Chromobacterium and Janthinobacterium

737

The Genera Phyllobacterium and Ochrobactrum

747

The Genus Derxia

751

The Genera Leptothrix and Sphaerotilus

758

The Lithoautotrophic Ammonia-Oxidizing Bacteria hans-peter koops, ulrike purkhold, andreas pommerening-röser,

778

3.2.3 3.2.4 3.2.5 3.2.6 3.2.7 3.2.8 3.2.9 3.2.10 3.2.11 3.2.12

alison weiss

noel r. krieg

bruno pot, monique gillis and jozef de ley anne willems and paul de vos monique gillis and jozef de ley

jean swings, bart lambert, karel kersters and barry holmes jan hendrick becking stefan spring

gabriele timmermann and michael wagner

Contents

xxi

The Genus Thiobacillus

812

The Genera Simonsiella and Alysiella brian p. hedlund and daisy a. kuhn

828

Eikenella corrodens and Closely Related Bacteria

840

The Genus Burkholderia

848

3.2.17

The Nitrite-Oxidizing Bacteria

861

3.2.18

The Genera Azoarcus, Azovibrio, Azospira and Azonexus barbara reinhold-hurek and thomas hurek

873

3.2.13 3.2.14 3.2.15 3.2.16

lesley a. robertson and j. gijs kuenen

edward j. bottone and paul a. granato donald e. woods and pamela a. sokol aharon abeliovich

893

Index

Volume 6 3.

Proteobacteria

3.3.

Gamma Subclass

3.3.1

New Members of the Family Enterobacteriaceae j. michael janda

3.3.2

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

The Genus Edwardsiella

72

The Genus Citrobacter

90

The Genus Shigella

99

3.3.3 3.3.4 3.3.5 3.3.6 3.3.7 3.3.8

rodney a. welch

sharon l. abbott and j. michael janda diana borenshtein and david b. schauer yves germani and philippe j. sansonetti

5

The Genus Salmonella

123

The Genus Klebsiella

159

craig d. ellermeier and james m. slauch sylvain brisse, francine grimont and patrick a. d. grimont

xxii

Contents

The Genus Enterobacter

197

The Genus Hafnia

215

The Genus Serratia

219

3.3.12

The Genera Proteus, Providencia, and Morganella

245

3.3.13

Y. enterocolitica and Y. pseudotuberculosis elisabeth carniel, ingo autenrieth, guy cornelis, hiroshi fukushima, françoise guinet, ralph isberg, jeannette pham, michael prentice, michel simonet,

270

3.3.14

Yersinia pestis and Bubonic Plague

399

3.3.15

Erwinia and Related Genera clarence i. kado

443

The Genera Photorhabdus and Xenorhabdus

451

3.3.17

The Family Vibrionaceae j. j. farmer, iii

495

3.3.18

The Genera Vibrio and Photobacterium j. j. farmer, iii and f. w. hickman-brenner

508

The Genera Aeromonas and Plesiomonas

564

The Genus Alteromonas and Related Proteobacteria valery v. mikhailov, lyudmila a. romanenko and elena p. ivanova

597

Nonmedical: Pseudomonas

646

3.3.22

Pseudomonas aeruginosa timothy l. yahr and matthew r. parsek

704

3.3.23

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

714

Xylophilus

741

3.3.9 3.3.10 3.3.11

francine grimont and patrick a. d. grimont megan e. mcbee and david b. schauer francine grimont and patrick a. d. grimont jim manos and robert belas

mikael skurnik and georges wauters

3.3.16

3.3.19 3.3.20

3.3.21

robert brubaker

noel boemare and raymond akhurst

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

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

nickolas panopoulos 3.3.24

anne willems and monique gillis

Contents

xxiii

The Genus Acinetobacter

746

3.3.26

The Family Azotobacteraceae

759

3.3.27

The Genera Beggiatoa and Thioploca andreas teske and douglas c. nelson

784

3.3.28

The Family Halomonadaceae david r. arahal and antonio ventosa

811

The Genus Deleya

836

The Genus Frateuria

844

3.3.31

The Chromatiaceae johannes f. imhoff

846

3.3.32

The Family Ectothiorhodospiraceae johannes f. imhoff

874

3.3.33

Oceanospirillum and Related Genera josé m. gonzález and william b. whitman

887

3.3.34

Serpens flexibilis: An Unusually Flexible Bacterium robert b. hespell

916

The Genus Psychrobacter

920

The Genus Leucothrix

931

The Genus Lysobacter

939

The Genus Moraxella

958

Legionella Species and Legionnaire’s Disease

988

3.3.25

3.3.29 3.3.30

3.3.35 3.3.36 3.3.37 3.3.38 3.3.39 3.3.40 3.3.41 3.3.42

kevin towner

jan hendrick becking

karel kersters jean swings

john p. bowman

thomas d. brock

hans reichenbach john p. hays

paul h. edelstein and nicholas p. cianciotto The Genus Haemophilus

1034

The Genus Pasteurella

1062

The Genus Cardiobacterium

1091

doran l. fink and joseph w. st. geme, iii henrik christensen and magne bisgaard sydney m. harvey and james r. greenwood

xxiv

3.3.43 3.3.44 3.3.45 3.3.46

3.3.47

Contents

The Genus Actinobacillus

1094

The Genus Francisella

1119

Ecophysiology of the Genus Shewanella kenneth h. nealson and james scott

1133

The Genus Nevskia

1152

The Genus Thiomargarita

1156

janet i. macinnes and edward t. lally francis nano and karen elkins

heribert cypionka, hans-dietrich babenzien, frank oliver glöckner and rudolf amann heide n. schulz

1165

Index

Volume 7 3.

Proteobacteria

3.4

Delta Subclass

3.4.1

The Genus Pelobacter

bernhard schink

5

The Genus Bdellovibrio

12

3.4.3

The Myxobacteria lawrence j. shimkets, martin dworkin and hans reichenbach

31

3.5.

Epsilon Subclass

3.4.2

3.5.1 3.5.2 3.5.3

edouard jurkevitch

The Genus Campylobacter

119

The Genus Helicobacter

139

The Genus Wolinella

178

trudy m. wassenaar and diane g. newell jay v. solnick, jani l. o’rourke, peter van damme and adrian lee jörg simon, roland gross, oliver klimmek and achim kröger

4.

Spirochetes

4.1

Free-Living Saccharolytic Spirochetes: The Genus Spirochaeta susan leschine, bruce j. paster and ercole canale-parola

195

Contents

4.2

4.3 4.4 4.5 4.6

xxv

The Genus Treponema

211

The Genus Borrelia

235

The Genus Leptospira

294

Termite Gut Spirochetes john a. breznak and jared r. leadbetter

318

The Genus Brachyspira

330

steven j. norris, bruce j. paster, annette moter and ulf b. göbel melissa j. caimano ben adler and solly faine

thaddeus b. stanton

5.

Chlorobiaceae

5.1

The Family Chlorobiaceae

6.

Bacteroides and Cytophaga Group

jörg overmann

359

The Medically Important Bacteroides spp. in Health and Disease

381

The Genus Porphyromonas

428

An Introduction to the Family Flavobacteriaceae

455

The Genus Flavobacterium

481

The Genera Bergeyella and Weeksella

532

6.6

The Genera Flavobacterium, Sphingobacterium and Weeksella

539

6.7

The Order Cytophagales

549

The Genus Saprospira

591

The Genus Haliscomenobacter

602

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

605

6.1 6.2 6.3 6.4 6.5

6.8 6.9 6.10

c. jeffrey smith, edson r. rocha and bruce j. paster frank c. gibson and caroline attardo genco jean-françois bernardet and yasuyoshi nakagawa jean-françois bernardet and john p. bowman celia j. hugo, brita bruun and piet j. jooste barry holmes

hans reichenbach hans reichenbach

eppe gerke mulder and maria h. deinema

xxvi

6.11 6.12 6.13

6.14

6.15 6.16

Contents

The Genera Empedobacter and Myroides celia j. hugo, brita bruun and piet j. jooste

630

The Genera Chryseobacterium and Elizabethkingia

638

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

The Genera Rhodothermus, Thermonema, Hymenobacter and Salinibacter

712

e. r. leadbetter

aharon oren

7. 7.1

677

Chlamydia The Genus Chlamydia—Medical

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,

741

757

heinz schlesner and erko stackebrandt

9. 9.1

Thermus The Genus Thermus and Relatives

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

797

10.

Chloroflexaceae and Related Bacteria

10.1

The Family Chloroflexaceae

815

The Genus Thermoleophilum

843

The Genus Thermomicrobium

849

10.2 10.3

satoshi hanada and beverly k. pierson jerome j. perry jerome j. perry

Contents

10.4

The Genus Herpetosiphon

natuschka lee and hans reichenbach

xxvii

854

11.

Verrucomicrobium

11.1

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

12.

Thermotogales

12.1

Thermotogales

13.

Aquificales

13.1

Aquificales

14.

Phylogenetically Unaffiliated Bacteria

14.1

Morphologically Conspicuous Sulfur-Oxidizing Eubacteria jan w. m. la rivière and karin schmidt

941

The Genus Propionigenium

955

The Genus Zoogloea

960

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

971

14.2 14.3 14.4

robert huber and michael hannig

robert huber and wolfgang eder

bernhard schink

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

lynn margulis and gregory hinkle

14.5 14.6 14.7 14.8 14.9

881

899

925

Streptobacillus moniliformis james r. greenwood and sydney m. harvey

983

The Genus Toxothrix

986

The Genus Gallionella

990

The Genera Caulococcus and Kusnezovia jean m. schmidt and georgi a. zavarzin

996

The Genus Brachyarcus

998

peter hirsch

hans h. hanert

peter hirsch

xxviii

14.10 14.11

14.12 14.13 Index

Contents

The Genus Pelosigma

1001

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

1005

The Genus Fusobacterium

1016

Prokaryotic Symbionts of Amoebae and Flagellates

1028

peter hirsch

tor hofstad

kwang w. jeon

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

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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

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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

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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

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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

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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

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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

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

Francis Nano Department of Biochemistry & Microbiology University of Victoria Victoria V8W 3PG Canada

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

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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

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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

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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

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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

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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

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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

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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

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Prokaryotes (2006) 2:3–15 DOI: 10.1007/0-387-30742-7_1

CHAPTER 1.1 c i notkna lP

sus reV

e l i s seS

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fo

se toyrakorP

Planktonic Versus Sessile Life of Prokaryotes KEVIN C. MARSHALL

Because of the extremely small size of most prokaryotic organisms, the limits on what is meant by the terms planktonic and sessile require definition. According to the Oxford English Dictionary, planktonic refers to “drifting or floating organic life found at various depths in the ocean or fresh water.” At the micrometer level, a planktonic habitat for prokaryotes can also encompass water films around soil particles, saliva in the mouth, fluids in the intestinal lumen, serum in blood vessels, and urine in the bladder and urinary tract. Sessile, on the other hand, means “immediately attached, without a footstalk.” Again, one can extend this definition to include those prokaryotes directly adhering to surfaces, those attaching by means of a holdfast at the end of a prostheca (e.g., Caulobacter), those embedded in biofilms developing as a result of extracellular polymer production by bacteria colonizing surfaces, and those colonizing mucus excreted by higher organisms (as in the gastrointestinal tract and the mucigel of plant roots). Most microbiologists, oriented by their training to the study of pure cultures, regard suspension culture as the normal state for growth of these organisms. This is particularly true for research into the physiology and biochemistry of bacteria, whereby homogeneous suspensions of bacteria are readily harvested and manipulated for experimental purposes. The reality of prokaryotic life in natural habitats is that many organisms spend part or all of their life spans attached to surfaces (Marshall, 1976). However, recently there has been a veritable explosion in research devoted to understanding the behavior of bacteria at surfaces (Beachey, 1980; Bitton and Marshall, 1980; Marshall, 1984; Savage and Fletcher, 1985). Many questions arise regarding the association of bacteria with surfaces. It is my aim in this chapter to consider the current state of knowledge concerning the following questions: How do prokaryotes adhere to surfaces? Is there a single, all-embracing mechanism or a range of mechanisms of adhesion in different organisms? Are some prokaryotes especially adapted to a sessile existence? Are particular organisms homoge-

neous in their adhesive characteristics or are they variable in their response to surfaces? Once attached to a surface, do prokaryotes always remain in a sessile state or do they return to the planktonic state at some stage? Do prokaryotes gain any real advantage from being associated with surfaces? Are certain prokaryotes specifically adapted to the colonization of excreted mucous layers? Are sessile bacteria in a different physiological state from planktonic organisms; that is, do prokaryotes exhibit a physiological response to contact with a surface? If they show such responses, what physicochemical factors are responsible for inducing the responses?

Mechanisms of Adhesion to Surfaces Full details of proposed mechanisms of adhesion of prokaryotes to solid surfaces have been presented elsewhere (Marshall, 1985, 1986a) so only a brief outline will be presented in this paper.

Transport Processes Water currents induced by temperature and gravity (fluid dynamic forces) provide the major mechanism for the transport of planktonic bacteria over large distances. When bacteria and other particles in flowing water are transported to the region of the boundary layer near a solid surface, a lift force directs the bacteria toward the surface where fluid frictional forces slow them down (Characklis, 1981a) and deposit them in the vicinity of the surface. Sedimentation is of significance only when bacteria are aggregated together or are attached to particles. Individual bacteria behave essentially as colloidal particles (Marshall, 1976) and tend to remain in suspension. Nutrient gradients may become established across the boundary layer near some surfaces and these may provide opportunities for chemotactic responses towards the surfaces by motile bacteria. Brownian motion can account for random movement of very small bacteria within the quiescent water of a boundary layer near a surface (Marshall, 1976).

4

K.C. Marshall

Long-Range Forces Bacteria in the vicinity of a solid-liquid interface frequently show an instantaneous but reversible attraction to the interface (Marshall et al., 1971a), and an attempt has been made to explain this reversible attraction by means of the colloid stability (DLVO named for the originators— Derjaguin, Landau, Verwey, and Overbeek) theory. That this attraction is reversible is shown by the fact that the bacteria can be removed from the solid surface by gentle shearing forces. The DLVO theory accounts, at least in part, for the attraction of a negatively charged bacterium to a negatively charged substratum surface at the “secondary attraction minimum” resulting from the interaction between London-van der Waals attraction forces and electrical repulsion forces in the overlapping double layers of cations surrounding the negatively charged surfaces. In terms of the DLVO theory, a bacterial cell would be held at a distance of some 10 nm from the surface by repulsion forces. Problems in applying DLVO theory to biological systems have been raised by Pethica (1980) and Rutter and Vincent (1980), especially when the complexity of the bacterial cell envelope and the extracellular components are taken into account. However, Busscher and Weerkamp (1987) have argued strongly in favor of such long-range forces in the initial attraction of bacteria to surfaces.

Short-Range Forces Certain bacteria irreversibly attach to surfaces very rapidly (Fletcher, 1980), whereas other bacteria require a significant time of exposure to the surface before becoming firmly attached (Marshall et al., 1971a). Irreversible attraction is shown by the fact that the bacteria cannot be removed by moderate shear forces. What is the mechanism of this firm adhesion of bacteria to surfaces? Early observations indicated that polymer bridging by extracellular components of cells to the substratum surface (Fig. 1) resulted

CHAPTER 1.1

in firm adhesion (Marshall and Cruickshank, 1973; Fletcher and Floodgate, 1973), and these observations have been confirmed for many systems (Corpe, 1980; Costerton et al., 1981). These extracellular polymers have a small radius of curvature and can overcome any repulsion barrier near a surface and, thus, can bind the cell to the surface using a variety of short-range forces. These forces include: 1) chemical bonds (electrostatic, covalent, and hydrogen); 2) dipole interactions (dipole-dipole, dipole-induced dipole, and ion-dipole); and 3) hydrophobic interactions (Rutter and Vincent, 1980). Adhesion to surfaces in nature is generally considered to be nonspecific. That is, the bacteria adhere to a wide variety of different inanimate, and possibly animate, surfaces with varying degrees of adhesive strength. Bridging polymers involved in most cases of nonspecific adhesion are either extracellular polysaccharides, proteins, or glycoproteins. The precise mechanisms whereby such polymers interact with a range of substratum surfaces is not known, but it almost certainly involves various combinations of the short-range forces listed above. Specific adhesion involves lectin-receptortype mechanisms, in which a proteinaceous substance (lectin) on the bacterial surface reacts with a complementary carbohydrate receptor on another cell type (Switalski et al., 1989). The best-described examples of specific adhesion involve the attachment of pathogenic bacteria to the host cell surfaces they infect. However, specific attachment of bacteria to the heterocysts of the cyanobacterium Anabaena has been described (Lupton and Marshall, 1981).

Thermodynamic Approach to Bacterial Adhesion Various workers have attempted to relate the extent of bacterial adhesion to the variation in surface free energy of the substratum, with very variable results (Dexter et al., 1975; Fletcher and Loeb, 1979). More detailed studies revealed that, in addition to the substratum-surface free energy, it was necessary to consider the bacterium-surface free energy and the surface tension of the liquid (Absolom et al., 1983; Pringle and Fletcher, 1983). The change in free energy associated with bacterial adhesion (DFadh) is given by: DFadh = g BS - g BL - g SL

1 µm Fig. 1. Perpendicular adhesion of a marine bacterium to a solid plastic surface. The extracellular polymeric substances bridging between the cell and the surface are present only at the adhesive pole of the cell. (Courtesy of R. H. Cruickshank.)

where gBS, gBL, and gSL are the bacteriumsubstratum, bacterium-liquid, and substratumliquid interfacial tensions, respectively. Bacterial adhension is favored if the process results in a free energy decrease. In general, Absolom et al. (1983) found good agreement between bacterial adhesion to a variety of substrata and the adhe-

CHAPTER 1.1

Fig. 2. Colonization of a glass surface, rendered hydrophobic by treatment with silane, by a marine bacterium after 16 h exposure. The condensed extracellular polymeric substances are clearly visible, as a result of drying on a cold stage. (Courtesy of T. Neu.)

sion behavior predicted by the thermodynamic model.

Detachment of Bacteria from Surfaces Not all cells remain adherent at the surface. Mechanisms of detachment include fluid shear forces (Marshall et al., 1971a), changes in surface free energy of the substratum (Busscher et al., 1986) or the organism (Rosenberg et al., 1983; Fattom and Shilo, 1984), reproductive mechanisms (Power and Marshall, 1988), and enzymatic degradation of adhesive structures. In most cases, however, the majority of adhering bacteria remain at the surface, where they are capable of growth, reproduction (Fig. 2) (Lawrence and Caldwell, 1987; Power and Marshall, 1988; Szewzyk and Schink, 1988), and even biofilm formation. A biofilm consists of cells immobilized at a substratum surface and frequently embedded in an organic polymer matrix of microbial origin (Characklis and Marshall, 1990). Other practical aspects of bacterial detachment from surfaces will be considered in later sections.

Occurrence of Sessile Prokaryotes Microbial Succession at Surfaces Early reports indicated that very small bacteria were the primary colonizers of surfaces immersed in seawater and were succeeded by conventional rod-shaped and, somewhat later, by prosthecate bacteria (Marshall et al., 1971b). It was realized that the initial colonizing organisms were starvation-survival forms (Morita, 1982) that eventually produced cellular growth at surfaces and thus gave rise to rod-shaped

Planktonic Versus Sessile Life of Prokaryotes

5

forms (Dawson et al., 1981; Power and Marshall, 1988). Early colonizing organisms tend to be Gram-negative bacteria, particularly species of Pseudomonas, Flavobacterium, and Achromobacter, followed later by prosthecate bacteria (Corpe, 1973). Gram-positive bacteria have rarely been recorded on surfaces in aquatic habitats, although there have been recent reports of significant numbers of Gram-positive bacteria on surfaces associated with groundwater (KölbelBoelke and Hirsch, 1989) and on the seagrass Zostera capricorni (Angles, 1988). The numbers, overall biomass, and diversity of attached microorganisms increased with increasing time of immersion of a surface (Jordon and Staley, 1976). Scanning electron microscopic studies also have revealed a progression from rod-shaped primary colonizers, to prosthecate forms, and then to a complex biofilm whose composition varies with the nature of the exposed surface and with time (Gerchakov et al., 1977; Marszalek et al., 1979; Dempsey, 1981). Even in illuminated waters, microalgae are not primary colonizers of surfaces (Marshall et al., 1971b; Corpe, 1973; Jordon and Staley, 1976), but extensive development of diatoms, fungi, and protozoa has been observed following bacterial biofilm formation (Gerchakov et al., 1977; Marszalek et al., 1979). Biologically inert substrata, such as stainless steel or glass, were colonized rapidly following immersion in seawater and produced a complex, two-tiered, microfouling layer (Gerchakov et al., 1977; Marszalek et al., 1979; Dempsey, 1981). The first stage of colonization consisted mainly of bacteria followed by nonmotile diatoms and fungi, whereas the second stage, which appeared after a 5-week exposure, consisted of large, colonial, motile diatoms, other diatoms, flagellates, and ciliates. On the other hand inhibitory substrata, such as copper-nickel alloys or brass, were slowly fouled by bacteria capable of secreting mucoid extracellular polymeric substances (EPS). Such substrata eventually developed a much less diverse biofilm community than inert ones. Sequential establishment of sessile populations also occurs in freshwater streams (Geesey et al., 1977, 1978) and lakes (Paerl, 1980); in soils where the complexity and variability of the solid matrix makes adequate study difficult (Marshall, 1975; Stotzky, 1986); in the oral cavity (Bowden et al., 1979; Newman, 1980); in the gastrointestinal tract, where the normal sessile biota plays an important role in preventing colonization by bacterial pathogens (Lee, 1980, 1985; Savage, 1980, 1984); and in the colonization of prosthetic devices employed in human patients (Gristina, 1987).

Biofilm Formation The combined effects of continuous adhesion and both growth and reproduction at surfaces

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K.C. Marshall

CHAPTER 1.1 Flow Sloughing Oxygen

Nutrients

Nitrate

AEROBIC

ANOXIC Predation Fermentation products

Cell

Iysis ANAEROBIC

A

MATURE

SO4=

S= SRB

BIOFILM

Fig. 3. Diagram of a section through a well-developed biofilm, showing bacteria embedded in an EPS matrix and the direction of decreasing gradients (arrows) of nutrients, oxygen, nitrate, and fermentation products. A predatory amoeba within the biofilm is shown at the left. SRB = sulfate reducing bacterium.

eventually gives rise to a macroscopic slime, or biofilm (Fig. 3). Biofilms are of considerable nuisance on artificial structures, such as ship hulls, hydroelectric pipelines, water reticulation systems, heat exchangers, oil rigs, and floating oceanographic equipment, but find useful applications in wastewater trickling-filter plants and other fixed-film systems, as well as in fluidized-bed fermenters. The development of a biofilm on a surface subjected to high shear rates may be described by a sigmoid-shaped curve, where the phase of biomass increase is a function of growth of attached bacteria along with further accretion of cells to the developing biofilm. The plateau of the curve represents the point at which the film penetrates the boundary (or viscous) sublayer (Characklis, 1981b). The final biofilm thickness is dependent on the magnitude of the fluid shear rate. Any protrusion of film irregularities above the viscous sublayer creates turbulence in the water flowing past the biofilm surface leading to frictional flow resistance. The colonization of mucous excreted by higher organisms (e.g.) the mucous blanket of the animal gastrointestinal tract (Lee, 1985); and the mucigel of plant roots (Rovira et al., 1979), leads to a partial or complete immobilization of cells in the mucous adjacent to the organism’s tissue. The final product in this instance bears a super-

ficial resemblance to a biofilm but its mode of origin is entirely different. Certain organisms, particularly spiral bacteria (Phillips and Lee, 1983), appear to have a selective advantage in penetrating and colonizing this viscous habitat.

Methods of Studying Sessile Prokaryotes Because of the inherent difficulty in directly observing the behavior of microorganisms at surfaces, a wide range of semidirect and indirect techniques have been employed to study adhesion, growth, biofilm development, and detachment from surfaces. Because of the different techniques needed for different surfaces and ecosystems, no attempt will be made here to give detailed instructions for the many techniques available but, rather, references to the descriptions of the original techniques will be provided.

Microscopy Many of the applications of various forms of microscopy in the study of sessile bacteria have been reviewed (Marshall, 1986b). Most studies involve the use of transmitted or incident light microscopy, or of transmission (TEM) or scanning (SEM) electron microscopy. For transmit-

CHAPTER 1.1

ted light microscopy, the use of transparent substrata (glass, mica, cellophane, polystyrene, etc.) as test surfaces is essential. Epifluorescence microscopy is necessary for translucent and opaque substrata (Zvyagintsev, 1962; Hobbie et al., 1977). Sessile bacteria may be observed by washing the exposed substratum to remove debris and loosely attached cells and then either staining, with conventional bacteriological stains or fluorescent dyes, or viewing directly with phase-contrast optics. The advantages and disadvantages of such techniques have been presented by Marshall (1986b). Novel techniques involving light microscopy include the use of submerged microscopy (Staley, 1971), capillary microscopy (Perfil’ev and Gabe, 1969), computer-enhanced image analysis (Caldwell and Germida, 1985), interference reflection microscopy (Fletcher, 1988), dialysis microculture (Duxbury, 1977), marked slides (Bott and Brock, 1970), soil films (Harris, 1972), transparent sections in tubular reactors to study biofilm development (Characklis, 1980), and light section microscopy to measure biofilm thickness (Loeb, 1980).

Other Methods of Study During the early stages of colonization of surfaces, and particularly if glass, plastic, metal, or wooden slides are immersed in an aqueous phase, bacteria adhering firmly to the surface may be cultured by washing the slides or coupons to remove loosely adhering organisms and then smearing the slide or coupon over the surface of a suitable agar plate (Marshall et al., 1971a). If a distinct biofilm has formed on a surface, the biofilm may be scraped from the surface, suspended in a suitable diluent, homogenized, a dilution series prepared, and aliquots of each dilution plated on an appropriate agar medium. Such methods suffer from the normal problems of selectiveness of the medium employed, and it is likely that some colonizing species (e.g., Caulobacter, Hyphomicrobium) are never obtained by such techniques. Often the use of special selective media is required in order to isolate particular organisms that may be obvious microscopically. In some cases, it may be necessary to resort to micromanipulation techniques to separate slow-growing or sensitive organisms from more aggressive or resistant species. The simple micromanipulation system devised by Skerman (1968) is especially recommended for this purpose. A variety of other methods have been adapted to estimate numbers of microorganisms or the total biomass found in a sessile state at surfaces. These include: measurement of radioactivity fol-

Planktonic Versus Sessile Life of Prokaryotes

7

lowing the uptake of labeled substrates (Brock, 1971; Lupton and Marshall, 1981), autoradiography (Fletcher, 1979; Bright and Fletcher, 1983), ATP determinations for total biomass (La Motta, 1976), muramic acid determinations for bacterial biomass (Moriarty, 1977), bacterial growth rates using thymidine incorporation (Moriarty, 1986), and determination of bacterial types at surfaces by phospholipid fatty acid signature analysis (Guckert et al., 1985) and by 16S rRNA sequence analysis (Pace et al., 1986; Weller and Ward, 1989). Other techniques that may prove valuable in analyzing biofilm composition and function include the use of Fourier transform infrared spectrophotometry (Nichols et al., 1985) and the use of microelectrodes to measure various gradients with depth of biofilms (Revsbech and Jørgensen, 1986) (Fig. 3).

Adaptation to the Sessile State Are certain prokaryotes uniquely adapted to a sessile form of life? The answer to this question is not simple because of the very wide range of bacteria that can be found on various surfaces. Several examples of different modes of sessile behavior will be considered in order to illustrate the complexity that may be encountered in natural habitats. Although many bacteria are capable of adhering to a wide variety of surfaces (nonspecific adhesion), the extent of adhesion on the various surfaces varies considerably. Some bacteria adhere best to hydrophobic surfaces (Fletcher and Loeb, 1979), some adhere best to hydrophilic surfaces (Dexter et al., 1975), whereas others adhere best to surfaces of more intermediate surface-free-energy values (Pringle and Fletcher, 1983). The conditions under which the bacteria are grown also modify the adhesive ability of various bacteria on a range of different surfaces (McEldowney and Fletcher, 1986). Many bacteria that require relatively high nutrient concentrations (copiotrophic bacteria) exist planktonically in oligotrophic waters in a state of starvation. These starvation-survival forms are characterized by a significant reduction in size and by lower endogenous respiration and heat output, and are often more adhesive than actively growing cells (Morita, 1982; Dawson et al., 1981; Humphrey and Marshall, 1984). Adhesion to surfaces by these starvationsurvival forms provides access to nutrients accumulated at the surfaces. The starved bacteria are able to scavenge these nutrients and metabolize them (Kefford et al. 1982; Kjelleberg et al., 1983), thereby leading to cellular growth and reproduction (Kjelleberg et al., 1982; Power and Marshall, 1988; Szewzyk and Schink, 1988). In

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K.C. Marshall

CHAPTER 1.1

Sessile

Planktonic

(a) motile daughter cell

(b)

detachment

(c) detachment capsule

(d)

division following drift from surface Fig. 4. Four mechanisms for alternating between the planktonic and sessile states: (a) a perpendicularly attached mother cell releases a motile daughter cell, as in Vibrio sp. DWI; (b) division of a cell adhering in a face-to-face manner, and release of a cell on utilization of a bound hydrophobic substrate, as in Pseudomonas sp. JD8; (c) detachment of a fimbrial-attached organism following the production of a hydrophilic capsule, as in Acinetobacter calcoaceticus; and (d) growth of a reversibly adhering organism at a surface and completion of the division phase following drift of the cell from the surface, as in Vibrio sp. MH3.

many marine environments, it appears that such small, starved bacteria are the primary colonizers of freshly immersed surfaces (Marshall et al., 1971b). Some copiotrophic bacteria seem unable to adhere firmly to surfaces, yet, under oligotrophic conditions, any starvation-survival forms approaching a surface are able to metabolize surfacebound substrates (Hermansson and Marshall, 1985) and exhibit both cellular growth and reproduction (Power and Marshall, 1988). Thus, nonadhesive bacteria do exist in the planktonic state but it is still possible for such organisms to benefit from association with surfaces. A particularly effective adaptation to the sessile state is the ability of many bacteria in nature to adhere in an orientation perpendicular to the surface (Fig. 4a; see also Fig. 1). Such prokaryotes appear to have either a specialized holdfast (Caulobacter) or a particularly adhesive

portion at one pole of the cell (Hyphomicrobium, Flexibacter, and Leucothrix). Such an orientation allows a very efficient contact both with the solid and the aqueous phases, as well as providing an effective means of releasing daughter cells into the planktonic state. An examination of this mode of orientation at solid surfaces revealed that both Hyphomicrobium and Flexibacter exhibited the same perpendicular orientation at air-water and oil-water interfaces (Marshall and Cruickshank, 1973). It was postulated that the pole of the cell approaching the interface was hydrophobic while the bulk of the cell was hydrophilic, and the hydrophobic pole was rejected from the water phase and aligned at the nonaqueous phase, regardless of whether it was solid, air, or oil (Marshall and Cruickshank, 1973). Some bacteria are adapted to growth at surfaces, yet possess various mechanisms to ensure that some cells return to the planktonic state. For instance, cells of the marine species Vibrio DW1 adhered to a surface in a perpendicular manner (Fig. 4a) and, following cellular growth of the starved cells to normal size, motile daughter cells were released at regular intervals (approximately 57 min) from the attached mother cells (Kjelleberg et al., 1982). Cells of the marine Pseudomonas sp. JD8 adhered in a face-to-face manner (Fig. 4b) and, following cellular growth and one division cycle the daughter cells slowly (about 0.15 mg/min) began to migrate away from each other while still adhering to the surface. After subsequent division cycles, similar migration patterns were observed but, eventually, some of the daughter cells detached from the surface (Power and Marshall, 1988). This slow migration was explained in terms of the cells being initially irreversibly attached to the hydrophobic stearic-acid-covered surface but, upon utilization of the fatty acid in the microenvironment around the cell, the cells became reversibly attached to the underlying hydrophilic substratum (Busscher et al., 1986) and were capable of some form of movement. As soon as the cells moved a short distance, however, they encountered more hydrophobic stearic acid and adhered irreversibly again until that substrate was utilized, and the cycle was repeated. When the bound substrate was essentially exhausted, cells detached from the underlying hydrophilic surface (Power and Marshall, 1988). Even the nonadhesive Vibrio MH3 (Fig. 4d) was able to grow from the small starvation-survival form to normal size and then begin the division cycle when exposed to surface-bound stearic acid (Power and Marshall, 1988). The dividing cells drifted away from the surface and completed the division cycle in the planktonic state. An interesting adaptation ensuring reversibility of the sessile state has been described in Acinetobacter calcoaceticus, which adheres

CHAPTER 1.1

reversibly to epithelial cells and oil by means of thin fimbriae (Fig. 4c). The adhesion of this bacterium is reversed as a result of the production of an excessive amount of extracellular emulsan that surrounds and thus masks the adhesive properties of the fimbriae (Rosenberg et al., 1983). Another example of reversible adhesion has been described in the cyanobacterium Phormidium, which in its sessile state possesses a hydrophobic surface but under certain conditions produces a hydrophilic capsule, thus allowing the organism to revert to the planktonic state (Fattom and Shilo, 1984). These studies emphasize the ability of some prokaryotes to take advantage of substrates adsorbed to surfaces, as well as revealing a variety of strategies for releasing daughter cells from the sessile to the planktonic state. As pointed out by Pedros-Alio and Brock (1983), a simple division into sessile and planktonic forms is overly simplistic. Different bacteria have a variety of mechanisms to attach at surfaces but they also possess a range of mechanisms for detachment in order to return to a planktonic existence.

Advantages of the Sessile State Nutrient Availability When a clean surface is immersed into a natural habitat, a molecular film rapidly forms on the surface as a result of adsorption of macromolecules and smaller hydrophobic molecules. This film serves to “condition” the surface, causing alterations in surface charge (Neihof and Loeb, 1974) and surface free energy (Baier, 1980). One of the most obvious advantages of the sessile state is the increased probability of access to nutrients accumulating at surfaces, particularly in flowing, oligotrophic conditions. ZoBell (1943) was the first to suggest that complex macromolecules adsorbed at surfaces would serve as concentrated sources of nutrients for organisms adhering at those surfaces. It was clearly demonstrated by Jannasch (1958) that the beneficial effect of surfaces in the presence of added complex nutrients only occurred at very low nutrient concentrations, where the level of nutrient in the aqueous phase was negligible and the nutrients had adsorbed to the surfaces. Many investigators comparing the activities of bacteria in the sessile and planktonic states have employed simple soluble substrates such as glucose and amino acids (Azam and Hodson, 1977; Berman, 1975; Berman and Stiller, 1977; Campbell and Baker, 1978; Ferguson and Palumbo, 1979; Fletcher, 1979, 1986; Hanson and Wiebe, 1977; Kirchman and Mitchell, 1982; Pedros-Alio and Brock, 1983; Riemann, 1978). In natural habitats, and particularly in low nutri-

Planktonic Versus Sessile Life of Prokaryotes

9

ent situations, such soluble substrates would be rapidly utilized by planktonic bacteria and would rarely encounter a substratum surface. Similarily, many of these low-molecular-weight substrates cannot adsorb to surfaces and would not be expected to concentrate these. If the substrates do adsorb, their availability for bacterial utilization is often reduced substantially (Gordon and Milero, 1985). In many field studies, filtration has been used to separate sessile and attached bacteria, but filtration can lead to problems in that: 1) shear forces involved in filtration are sufficient to remove some reversibly attached bacteria that are feeding at surfaces (Hermansson and Marshall, 1985); and 2) such reversibly attached bacteria may have fed, grown, and reproduced at the surface and then returned to the aqueous phase at some time prior to filtration (Power and Marshall, 1988). A more logical method of studying the activity of bacteria at surfaces is to provide substrates such as macromolecules or lower molecular weight hydrophobic molecules that are likely to adsorb at surfaces. Using surface-bound stearic acid as a model substrate, Kefford et al. (1982) and Kjelleberg et al. (1983) clearly demonstrated that a range of bacteria were capable of scavenging 14C-labeled stearic acid from a surface. In particular, a reversibly adhering Leptospira species rapidly utilized the labeled fatty acid, and 14 C-labeled bacteria were readily recovered from the planktonic state. A similar result was obtained with the nonadhesive marine Vibrio MH3 (Hermansson and Marshall, 1985), a result that emphasizes the fact that bacteria do not need to firmly adhere to surfaces in order to utilize substrates adsorbed at the surface. Subsequent studies have shown that starved bacteria adhering to surfaces where nutrients have accumulated not only metabolize the nutrients but are capable of cellular growth and reproduction (Kjelleberg et al., 1982; Power and Marshall, 1988; see also Fig. 4 a–d).

Protection from Harmful Factors Sessile bacteria appear to be more resistant to the inhibitory effects of antibacterial agents, such as antibiotics, chlorine, and heavy metals (Costerton et al., 1981). In relatively thick biofilms, this apparent resistance may be the result of the reaction of the agents with the outer layers of cells and, in the case of chlorine and heavy metals, reaction with the extracellular polymer that makes up the matrix of the biofilm. There is increasing evidence, however, that bacteria attached to surfaces are inherently more resistant to certain antibacterial agents than are planktonic forms, but the mechanism of this increased resistance is not understood. Bacteria below the biofilm-water interface are also pro-

10

K.C. Marshall

tected from external grazing by protozoa and metazoa. In addition, association of prokaryotes with various sizes of particles or colloidal clays can provide a degree of protection from parasitism by bacteriophage and Bdellovibrio, as well as from predation by amoebae and the lytic effects of certain gliding bacteria (Roper and Marshall, 1974, 1978).

Disadvantages of the Sessile State Sedimentation Although bacteria attached to particle surfaces may gain an advantage by utilization of adsorbed nutrients or by the dissolution of organic particles, such bacteria would sink to the sediments and would be unable to colonize new particle surfaces if mechanisms did not exist for their release or the release of daughter cells from the particle surfaces. As seen above, such mechanisms are common among sessile forms of bacteria (Fig. 4). It is precisely these phenomena of bacterial attachment, nutrient utilization, and recycling, and detachment that are continually occurring within “marine snow” in the pelagic zone of oceans (Alldredge, 1989).

Grazing Zooplankton are capable of ingesting planktonic bacteria but detritus feeders have been found to consume the bacteria growing on detritus particles rather than ingest the particles themselves (Fenchel and Jørgensen, 1977). Fenchel (1986) reported that the flagellate Bodo sp. spends about 45 sec ingesting a bacterium from a surface, during which time the flagellate does not move. Bodo normally slides over the substratum at a velocity of 3.5 mm/sec and only detects and ingests bacteria lying in a 1.0 mm wide band along the path of the flagellate. Zooplankton grazing on biofilm surfaces, however, may play a useful role in maintaining the bacteria near the biofilm surface in an active state of growth. Amoebae have been observed grazing well within the matrix of a biofilm (Mack et al., 1975) (see Fig. 3).

Gradients Decreasing gradients of nutrient and oxygen availability develop with increasing depth of a biofilm (Fig. 3) (Christensen and Characklis, 1990). Such gradients form as a result of diffusional resistance within the biofilm and of utilization of the nutrients and oxygen by microorganisms within the biofilm. Consequently, aerobic organisms near the biofilm-

CHAPTER 1.1

water-interface tend to be actively growing and create anoxic conditions at greater depths within the biofilm. If nitrate is present then some microorganisms at depth in the biofilm are capable of using the nitrate as an alternative to oxygen as an electron acceptor. Other aerobic organisms tend to be inactive, or even lyse, within the anoxic zone, whereas strict anaerobes and fermentative bacteria may be active in such sites. In biofilms developed on metallic surfaces, the activity of sulfate reducing bacteria (SRB) have been implicated in corrosion processes (Little et al., 1990).

Physiological Responses by Bacteria at Surfaces Observed Responses Probably the most obvious physiological response observed in bacteria associated with surfaces is cellular growth and, in some instances, reproduction (Jannasch, 1958; Bott and Brock, 1970; Kjelleberg et al., 1982; Pedros-Alio and Brock, 1983; Power and Marshall, 1988). Another possible response in bacteria to the physical presence of a surface is the timedependent appearance of firm adhesion, which may indicate the induction of suitable bridging polymer production by the surface-associated bacteria (Marshall et al., 1971a). The best documented response to a surface is the change observed in certain marine vibrios from a single, sheathed, polar flagellum in the planktonic stage to the production of multiple, lateral flagella when plated on an agar surface (Golten and Scheffers, 1975; de Boer et al., 1975; Belas and Colwell, 1982). Other reported responses include a reduction in size and an increase in endogenous respiration and in heat output by starving marine bacteria at interfaces in the absence of exogenous nutrients (Kjelleberg et al., 1982, 1983; Humphrey et al., 1983; Humphrey and Marshall, 1984). Also, attached bacteria show an increase in resistance to antibacterial substances (Costerton et al., 1981).

Control of Responses Silverman et al. (1984) have described two possible control mechanisms regulating bacterial responses at surfaces, namely, “responsive” and “variable” control (Fig. 5). Essentially, responsive control involves information processing, whereby the bacterium senses some environmental signal and responds accordingly. In the case of Vibrio parahaemolyticus, the response to a shift from an aqueous medium to an agar surface is to deregulate lateral flagella production

CHAPTER 1.1

(a)

Planktonic Versus Sessile Life of Prokaryotes

Responsive Control

Initial contact

(b)

11

Variable Control

Attach

Switch

Switch

Detach

Synthesize adhesin

Fig. 5. Strategies for responsive and for variable control of adhesive substance expression. (a) Responsive control, as shown by a shift from polar to lateral flagella in Vibrio parahaemolyticus. (b) Variable control, in which a fraction of the cells are preadapted to the fimbriated state, and attach to epithelial cells. Nonfimbriated variants detach and return to the aqueous phase. (From Silverman et al., 1984.)

(Fig. 5a). In the case of variable control, a fraction of the cells are preadapted, for example, to adhere to a particular surface, and individuals within the population are constantly switching among a variety of forms. For instance, a portion of the population may produce fimbriae and attach to epithelial cells (Fig. 5b). Nonadhesive variants of these cells arise and detach to return to the aqueous phase. Such phase variation in certain salmonellae results from a rearrangement of the DNA structure involving the inversion of part of the molecule containing a transcriptional control element.

Physicochemical Triggering of Responses Using lux gene fusion mutants, Belas et al. (1986) studied the responsive control of lateral gene expression when Vibrio parahaemolyticus was transferred from liquid to agar medium. They were able to show conclusively that the physicochemical factor triggering lateral flagella production was increased viscosity. Whether this surface effect was entirely the result of viscosity or whether it was also related to a reduction in water activity has not been tested.

Another important factor at surfaces that would result in metabolic, as well as cellular growth and reproduction responses, is the adsorption of organic nutrients at surfaces (Kefford et al., 1982; Kjelleberg et al., 1981; Hermansson and Marshall, 1985; Power and Marshall, 1988). Enhanced phosphorus uptake by attached bacteria has also been reported by Paerl and Merkel (1982). A further situation involving possible adsorption phenomena at surfaces is the finding by Humphrey and Marshall (1984) that changes in size, endogenous respiration, and heat output in starving marine bacteria at surfaces could be reproduced in the presence of surfactants and even when no surface was present. Many bacteria in nature produce surfactants, and these surfactants could adsorb to surfaces where they might trigger various responses in other bacteria adhering to the surfaces. Other possible explanations for the triggering of physiological responses in bacteria at surfaces include alterations in the proton motive force on the face of the cell nearest the surface (Ellwood et al., 1982) and possible cell deformation near a surface (Fletcher, 1984).

12

K.C. Marshall

Conclusions Although the sessile state is very common in bacteria in natural habitats, it is not a state limited to particular groups of organisms. All sessile bacteria are derived from the planktonic state and, in addition to active growth and metabolism at surfaces, these sessile organisms have also evolved a variety of methods to ensure that representatives of the population can return to the planktonic state. Such mechanisms include direct release of daughter cells, changes in the hydrophobicity of the sessile cells or of the substratum surface, exclusively reversible adhesion (subject to removal by gentle shear forces), and, possibly, enzymatic degradation of adhesive bridging polymers. Planktonic bacteria, on the other hand, possess a wide range of mechanisms whereby they can adhere to a variety of surfaces. In some instances these bacteria possess preformed adhesive polymers, whereas in other cases the bacteria appear to produce appropriate polymers following association with the surface. There is increasing evidence for responsive control of a number of physiological functions evident only at surfaces, but more detailed investigations are required to elucidate the nature of these physicochemical triggering mechanisms.

Literature Cited Absolom, D. R., F. V. Lamberti, Z. Policova, W. Zingg, C. J. van Oss, A. W. Neumann. 1983. Surface thermodynamics of bacterial adhesion. Appl. Environ. Microbiol. 46:90– 97. Alldredge, A. L. 1989. The significance of suspended detrital aggregates of marine snow as microhabitats in the pelagic zone of the ocean. 108–112. T. Hattori, Y. Ishida, Y. Maruyama, R. Y. Morita, and A. Uchida (ed.) Recent advances in microbial ecology. Japan Scientific Societies Press, Tokyo. Angles, M. L. 1988. Microbial colonization of Zostera capricorni in Botany Bay. B.Sc. (honors) thesis, University of New South Wales. Azam, F., R. E. Hodson. 1977. Size distribution and activity of marine microheterotrophs. Limnol. Oceanogr. 22:492–501. Baier, R. E. 1980. Substrate influence on adhesion of microorganisms and their resultant new surface properties. 59–104. G. Bitton and K. C. Marshall (ed.) Adsorption of microorganisms to surfaces. Wiley-Interscience, New York. Beachey, E. H. (ed.). 1980. Bacterial adherence. Chapman and Hall, London. Belas, M. R., R. R. Colwell. 1982. Adsorption kinetics of laterally and polarly flagellated Vibrio. J. Bacteriol. 151:1568–1580. Belas, R., M. Simon, M. Silverman. 1986. Regulation of lateral flagella gene transcription in Vibrio parahaemolyticus. J. Bacteriol. 167:210–218.

CHAPTER 1.1 Berman, T. 1975. Size fractionation of natural aquatic populations associated with autotrophic and heterotrophic carbon uptake. Mar. Biol. 33:215–220. Berman, T., M. Stiller. 1977. Simultaneous measurement of phosphorus and carbon uptake in Lake Kinneret by multiple isotopic labeling and differential filtration. Microb. Ecol. 3:279–288. Bitton, G., K. C. Marshall (ed.). 1980. Adsorption of microorganisms to surfaces. Wiley-Interscience, New York. Bott, T. L., T. D. Brock. 1970. Growth and metabolism of periphytic bacteria: Methodology. Limnol. Oceanogr. 15:333–342. Bowden, G. H. W., D. C. Ellwood, I. R. Hamilton. 1979. Microbial ecology of the oral cavity. Adv. Microb. Ecol. 3:135–217. Bright, J. J., M. Fletcher. 1983. Amino acid assimilation and electron transport system activity in attached and freeliving marine bacteria. Appl. Environ. Microbiol. 45:818–825. Brock, T. D. 1971. Microbial growth rates in nature. Bacteriol. Rev. 35:39–58. Busscher, H. J., M.H.M. J. C. Uyen, A. H. Weerkamp, A. H. Postma, J. Arends. 1986. Reversibility of adhesion of oral streptococci to solids. FEMS Microbiol. Lett. 35:303–306. Busscher, H. J., A. H. Weerkamp. 1987. Specific and nonspecific interactions in bacterial adhesion to solid substrata. FEMS Microbiol. Rev. 46:165–173. Caldwell, D. E., J. J. Germida. 1985. Evaluation of difference imagery for visualizing and quantitating microbial growth. Can. J. Microbiol. 31:35–44. Campbell, P. G. C., J. H. Baker. 1987. Estimation of bacterial production in freshwaters by the simultaneous measurement of [35S] sulfate and D-[3H] glucose uptake in the dark. Can. J. Microbiol. 24:939–946. Characklis, W. G. 1980. Biofilm development and destruction. Electric Power Research Institute. U.S. Report 902–1. Characklis, W. G. 1981a. Fouling biofilm development. A process analysis. Biotech. Bioeng. 23:1923–1960. Characklis, W. G. 1981b. Microbial fouling: a process analysis. 251–291. E. F. C. Somerscales and J. G. Knudsen (ed.) Fouling of heat transfer equipment.. Hemisphere Publ. Co. Washington, D.C. Characklis, W. G., K. C. Marshall. 1990. Biofilms: a basis for an interdisciplinary approach. 3–15. W. G. Characklis and K. C. Marshall (ed.) Biofilms. Wiley-Interscience, New York. Christensen, B. E., W. G. Characklis. 1990. Physical and chemical properties of biofilms. 93–130. W. G. Characklis and K. C. Marshall (ed.) Biofilms. Wiley-Interscience, New York. Corpe, W. A. 1973. Microfouling: the role of primary filmforming bacteria. 598–609. R. F. Acker, B. F. Brown, J. R. de Palma, and W. P. Iverson (ed.) Proc. 3rd Intern. Congr. Mar. Corrosion Fouling. Northwestern Univ. Press. Evanston, Illinois. Corpe, W. A. 1980. Microbial surface components involved in adsorption of microorganisms onto surfaces. 105–144. G. Bitton and K. C. Marshall (ed.) Adsorption of microorganisms to surfaces. Wiley-Interscience, New York. Costerton, J. W., R. J. Irvin, K. J. Cheng. 1981. The bacterial glycocalyx in nature and disease. Ann. Rev. Microbiol. 35:299–324.

CHAPTER 1.1 Dawson, M. P., B. A. Humphrey, K. C. Marshall. 1981. Adhesion, a tactic in the survival strategy of a marine vibrio during starvation. Curr. Microbiol. 6:195–198. de Boer, W. E., C. Golten, W. A. Scheffers. 1975. Effects of some physical factors on flagellation and swarming of Vibrio alginolyticus. Netherlands J. Sea Res. 9:197–213. Dempsey, M. J. 1981. Marine bacterial fouling: a scanning electron microscope study. Mar. Biol. 61:305–315. Dexter, S. C., J. D. Sullivan, Jr., J. Williams, III, S. W. Watson. 1975. Influence of substrate wettability on the attachment of marine bacteria to various surfaces. Appl. Microbiol. 30:298–308. Duxbury, T. 1977. A microperfusion chamber for studying the growth of bacterial cells. J. Appl. Bacteriol. 42:247– 251. Ellwood, D. C., C. W. Keevil, P. D. Marsh, C. M. Brown, J. N. Wardell. 1982. Surface associated growth. Phil. Trans. Roy. Soc. Lond. B297:517–532. Fattom, A., M. Shilo. 1984. Hydrophobicity as an adhesion mechanism of benthic cyanobacteria. Appl. Environ. Microbiol. 47:135–143. Fenchel, T. 1986. The ecology of heterotrophic microflagellates. Adv. Microb. Ecol. 9:57–97. Fenchel, T., B. B. Jørgensen. 1977. Detritus food chains of aquatic environments. Adv. Microb. Ecol. 1:1–58. Ferguson, R. L., A. V. Palumbo. 1979. Distribution of suspended bacteria in neritic waters south of Long Island during stratified conditions. Limnol. Oceanogr. 24:697– 705. Fletcher, M. 1979. A microautoradiographic study of the activity of attached and free-living bacteria. Arch. Microbiol. 122:271–274. Fletcher, M. 1980. The question of passive versus active attachment mechanisms in non-specific bacterial adhesion. 197–210. R. C. W. Berkeley, J. M. Lynch, J. Melling, P. R. Rutter, and B. Vincent (ed.) Microbial adhesion to surfaces. Ellis Horwood, Chichester. Fletcher, M. 1984. Comparative physiology of attached and free-living bacteria. 223–232. K. C. Marshall (ed.) Microbial adhesion and aggregation. Springer, Berlin. Fletcher, M. 1986. Measurement of glucose utilization by Pseudomonas fluorescens that are free living and that are attached to surfaces. Appl. Environ. Microbiol. 52:672–676. Fletcher, M. 1988. Attachment of Pseudomonas fluorescens to glass and influence of electrolytes on bacteriumsubstratum separation distance. J. Bacteriol. 170:2027– 2030. Fletcher, M., G. D. Floodgate. 1973. An electron microscopic demonstration of an acidic polysaccharide involved in the adhesion of a marine bacterium to solid surfaces. J. Gen. Microbiol. 74:325–334. Fletcher, M., G. I. Loeb. 1979. Influence of substratum characteristics on the attachment of a marine pseudomonad to solid surfaces. Appl. Environ. Microbiol. 37:67–72. Geesey, G. G., R. Mutch, J. W. Costerton, R. B. Green. 1978. Sessile bacteria: an important component of the microbial population in small mountain streams. Limnol. Oceanogr. 23:1214–1223. Geesey, G. G., W. T. Richardson, H. G. Yeomans, R. T. Irvin, J. W. Costerton. 1977. Microscopic examination of natural sessile bacterial populations from an alpine stream. Can. J. Microbiol. 23:1733–1736. Gerchakov, S. M., D. S. Marszalek, F. J. Roth, L. R. Udey. 1977. Succession of periphytic microorganisms on metal and glass surfaces. 203–211. V. Romanovsky (ed.) Proc.

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4th Intern. Congr. Mar. Corrosion Fouling, Centre de Recherches et d’Etudes Oceangraphiques. Boulogne, France. Golten, C., W. A. Scheffers. 1975. Marine vibrios isolated from water along the Dutch coast. Netherlands J. Sea Res. 9:351–364. Gordon, A. S., F. J. Milero. 1985. Adsorption mediated decrease in the biodegradation rate of organic compounds. Microb. Ecol. 11:289–298. Gristina, A. G. 1987. Biomaterial centred infection: Microbial adhesion versus tissue integration. Science 237: 1588–1595. Guckert, J. B., C. B. Antworth, P. D. Nichols, D. C. White. 1985. Phospholipid, ester-linked fatty acid profiles as reproducible assays for changes in prokaryotic community structure of estuarine sediments. FEMS Microbiol. Ecol. 31:147–158. Hanson, R. B., W. J. Wiebe. 1977. Heterotrophic activity associated with particulate size fractions in a Spartina alterniflora salt-marsh estuary, Sapelo Island, Georgia, U.S.A., and the continental shelf waters. Mar. Biol. 42:321–330. Harris, P. J. 1972. Micro-organisms in surface films from soil crumbs. Soil. Biol. Biochem. 4:105–106. Hermansson, M., K. C. Marshall. 1985. Utilization of surface localized substrate by non-adhesive marine bacteria. Microb. Ecol. 11:91–105. Hobbie, J. E., R. J. Daley, S. Jasper. 1977. Use of Nuclepore filters for counting bacteria by fluorescence microscopy. Appl. Environ. Microbiol. 33:1225–1228. Humphrey, B. A., S. Kjelleberg, K. C. Marshall. 1983. Responses of marine bacteria under starvation conditions at a solid-water interface. Appl. Environ. Microbiol. 45:43–47. Humphrey, B. A., K. C. Marshall. 1984. The triggering effect of surfaces and surfactants on heat output, oxygen consumption and size reduction of a starving marine Vibrio. Arch. Microbiol. 140:166–170. Jannasch, H. W. 1958. Studies on planktonic bacteria by means of a direct membrane filter method. J. Gen. Microbiol. 18:609–620. Jordan, T. L., J. T. Staley. 1976. Electron microscopic study of succession in the periphyton communities of Lake Washington. Microb. Ecol. 2:241–251. Kefford, B., S. Kjelleberg, K. C. Marshall. 1982. Bacterial scavenging: Utilization of fatty acids localized at a solidliquid interface. Arch. Microbiol. 133:257–260. Kirchman, D., R. Mitchell. 1982. Contribution of particlebound bacteria to total microheterotrophic activity in five ponds and two marshes. Appl. Environ. Microbiol. 43:200–209. Kjelleberg, S., B. A. Humphrey, K. C. Marshall. 1982. The effect of interfaces on small starved marine bacteria. Appl. Environ. Microbiol. 43:1166–1172. Kjelleberg, S., B. A. Humphrey, K. C. Marshall. 1983. Initial phases of starvation and activity of bacteria at surfaces. Appl. Environ. Microbiol. 46:978–984. Kölbel-Boelke, J., P. Hirsch. 1989. Comparative physiology of biofilm and suspended organisms in the groundwater environment. 221–238. W. G. Characklis and P. A. Wilderer (ed.) Structure and function of biofilms. Dahlem Konferenzen, John Wiley and Sons, New York. La Motta, E. J. 1976. Kinetics of growth and substrate uptake in a biological film system. Appl. Environ. Microbiol. 31:286–293. Lawrence, J. R., D. E. Caldwell. 1987. Behavior of bacterial stream populations within the hydrodynamic boundary

14

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layers of surface microenvironments. Microb. Ecol. 14:15–27. Lee, A. 1980. Normal flora of animal intestinal surfaces. 145– 173. G. Bitton and K. C. Marshall (ed.) Adsorption of microorganisms to surfaces. Wiley-Interscience, New York. Lee, A. 1985. Neglected niches: The microbial ecology of the gastrointestinal tract. Adv. Microb. Ecol. 8:115–162. Little, B. J., P. A. Wagner, W. G. Characklis, W. Lee. 1990. Microbial corrosion. 635–670. W. G. Characklis and K. C. Marshall (ed.) Biofilms. Wiley-Interscience, New York. Loeb, G. I. 1980. Measurement of microbial marine fouling films by light section microscopy. Mar. Technol. Soc. J. 14:17–30. Lupton, F. S., K. C. Marshall. 1981. Specific adhesion of bacteria to heterocysts of Anabaena spp. and its ecological significance. Appl. Environ. Microbiol. 42:1085–1092. Mack, W. N., J. P. Mack, A. O. Ackerson. 1975. Microbial film development in a trickling filter. Microb. Ecol. 2:215– 226. Marshall, K. C. 1975. Clay mineralogy in relation to survival of soil bacteria. Ann. Rev. Phytopath. 13:357–373. Marshall, K. C. 1976. Interfaces in microbial ecology. Harvard University Press. Cambridge, MA. Marshall, K. C. (ed.) 1984. Microbial adhesion and aggregation. Springer, Berlin. Marshall, K. C. 1985. Mechanisms of bacterial adhesion at solid-water interfaces. 131–161. D. C. Savage and M. Fletcher (ed.) Bacterial adhesion: Mechanisms and physiological significance. Plenum Press, New York. Marshall, K. C. 1986a. Adsorption and adhesion processes in microbial growth at interfaces. Adv. Colloid. Interface. Sci. 25:59–86. Marshall, K. C. 1986b. Microscopic methods for the study of bacterial behavior at inert surfaces. J. Microbiol. Methods 4:217–227. Marshall, K. C., R. H. Cruickshank. 1973. Cell surface hydrophobicity and the orientation of certain bacteria at interfaces. Arch. Mikrobiol. 91:29–40. Marshall, K. C., R. Stout, R. Mitchell. 1971a. Mechanism of the initial events in the sorption of marine bacteria to surfaces. J. Gen. Microbiol. 68:337–348. Marshall, K. C., Stout, R., Mitchell, R. 1971b. Selective sorption of bacteria from seawater. Can. J. Microbiol. 17:1413–1416. Marszalek, D. S., S. M. Gerchakov, L. R. Udey. 1979. Influence of substrate composition on marine microfouling. Appl. Environ. Microbiol. 38:987–995. McEldowney, S., M. Fletcher. 1986. Effect of growth conditions and surface characteristics of aquatic bacteria on their attachment to solid surfaces. J. Gen. Microbiol. 132:513–523. Moriarty, D. J. W. 1977. Improved method using muramic acid to estimate biomass of bacteria in sediments. Oecolgia 26:317–323. Moriarty, D. J. W. 1986. Measurement of bacterial growth rates in aquatic systems from rates of nucleic acid synthesis. Adv. Microb. Ecol. 9:245–292. Morita, R. Y. 1982. Starvation-survival of heterotrophs in the marine environment. Adv. Microb. Ecol. 6:171–198. Neihof, R., G. Loeb. 1974. Dissolved organic matter in seawater and the electric charge of immersed surfaces. J. Mar. Res. 32:5–12. Newman, H. N. 1980. Retention of bacteria on oral surfaces. 207–251. G. Bitton and K. C. Marshall (ed.) Adsorption

CHAPTER 1.1 of microorganisms to surfaces. Wiley-Interscience, New York. Nichols, P. D., J. M. Henson, J. B. Guckert, D. E. Nivens, D. C. White. 1985. Fourier transform-infrared spectroscopic methods for microbial ecology: Analysis of bacteria, bacteria polymer mixtures and biofilms. J. Microbiol. Methods 4:79–94. Pace, N. R., D. A. Stahl, D. J. Lane, G. J. Olsen. 1986. The analysis of natural microbial populations by ribosomal RNA sequences. Adv. Microb. Ecol. 9:1–55. Paerl, H. W. 1980. Attachment of microorganisms to living and detrital surfaces in freshwater systems. 375–402. G. Bitton and K. C. Marshall (ed.) Adsorption of microorganisms to surfaces. Wiley-Interscience, New York. Paerl, H. W., S. M. Merkel. 1982. Differential phosphorus assimilation in attached vs. unattached microorganisms. Arch. Hydrobiol. 93:125–134. Pedros-Alio, C., T. D. Brock. 1983. The importance of attachment to particles for planktonic bacteria. Arch. Hydrol. 98:354–379. Perfil’ev, B. V., D. R. Gabe. 1969. Capillary methods of investigating micro-organisms (translated from Russian by J. M. Shewan). Univ. of Toronto Press, Toronto. Pethica, B. A. 1980. Microbial and cell adhesion. 19–45. R. C. W. Berkeley, J. M. Lynch, J. Melling, P. R. Rutter, and B. Vincent (ed.) Microbial adhesion to surfaces. Ellis Horwood, Chichester. Phillips, M. W., A. Lee. 1983. Isolation and characterization of a spiral bacterium from the crypts of rodent gastrointestinal tracts. Appl. Environ. Microbiol. 45:675– 683. Power, K., K. C. Marshall. 1988. Cellular growth and reproduction of marine bacteria on surface-bound substrate. Biofouling 1:163–174. Pringle, J. H., M. Fletcher. 1983. Influence of substratum wettability on attachment of freshwater bacteria to solid surfaces. Appl. Environ. Microbiol. 45:811–817. Revsbech, N. P., B. B. Jørgensen. 1986. Microelectrodes: their use in microbial ecology. Adv. Microb. Ecol. 9:252–293. Riemann, B. 1978. Differentiation between heterotrophic and photosynthetic plankton by size fractionation, glucose uptake, ATP, and chlorophyll content. Oikos 31:358–367. Roper, M. M., K. C. Marshall. 1974. Modification of the interaction between Escherichia coli and bacteriophage in saline sediment. Microb. Ecol. 1:1–14. Roper, M. M., K. C. Marshall. 1978. Effects of a clay mineral on microbial predation and parasitism on Escherichia coli. Microb. Ecol. 4:279–289. Rosenberg, E., A. Gottlieb, M. Rosenberg. 1983. Inhibition of bacterial adherence to epithelial cells and hydrocarbons by emulsan. Infect. Immun. 39:1024–1028. Rovira, A. D., R. D. Foster, J. K. Martin. 1979. Note on terminology: Origin, nature and nomeclature of the organic materials in the rhizosphere. J. L. Harley and R. S. Russell (ed.) The Soil Root Interface. Academic Press, London, 1–4. Rutter, P. R., B. Vincent. 1980. The adhesion of microorganisms to surfaces: physico-chemical aspects. 79–93. R. C. W. Berkeley, J. M. Lynch, J. Melling, P. R. Rutter and B. Vincent (ed.) Microbial adhesion to surfaces. Ellis Horwood, Chichester. Savage, D. C. 1980. Colonization by and survival of pathogenic bacteria on intestinal mucosal surfaces. 175–206. G. Bitton and K. C. Marshall (ed.) Adsorption of microorganisms to surfaces. Wiley-Interscience, New York.

CHAPTER 1.1 Savage, D. C. 1984. Activities of microorganisms attached to living surfaces. 233–249. K. C. Marshall (ed.) Microbial adhesion and aggregation. Dahlem Konferenzen, Springer, Berlin. Savage, D. C., M. M. Fletcher (ed.). 1985. Bacterial adhesion: Mechanisms and physiological significance. Plenum Press, New York. Silverman, M., R. Belas, M. Simon. 1984. Genetic control of bacterial adhesion. 95–107. K. C. Marshall (ed.) Microbial adhesion and aggregation. Springer, Berlin. Skerman, V. B. D. 1968. A new type of micromanipulator and microforge. J. Gen. Microbiol. 54:287–297. Staley, J. T. 1971. Growth rates of algae determined in situ using an immersed microscope. J. Phycol. 7:13–17. Stotzky, G. 1986. Influence of soil mineral colloids on metabolic processes, growth, adhesion, and ecology of microbes and viruses. 305–428. Interactions of soil min-

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erals with natural organics and microbes. Soil Science Society of America Special Publication No. 17. Madison, Wisconsin. Switalski, L., M. Höök, E. Beachey (ed.). 1989. Molecular mechanisms of microbial adhesion. Springer, New York. Szewzyk, U., B. Schink. 1988. Surface colonization by and life cycle of Pelobacter acidigallici studied in a continuous flow microchamber. J. Gen. Microbiol. 134:183– 190. Weller, R., D. M. Ward. 1989. Selective recovery of 16SrRNA sequences from natural microbial communities in the form of cDNA. Appl. Environ. Microbiol. 55:1818–1822. ZoBell, C. E. 1943. The effect of solid surfaces upon bacterial activity. J. Bacteriol. 46:39–56. Zvyagintsev, D. G. 1962. Adsorption of microorganisms by soil particles. Soviet. Soil. Sci. 140–144.

Prokaryotes (2006) 2:16–31 DOI: 10.1007/0-387-30742-7_2

CHAPTER 1.2 l a i re t caB

no i sehdA

Bacterial Adhesion ITZHAK OFEK, NATHAN SHARON AND SOMAN N. ABRAHAM

Introduction It is now well established that to initiate infection at a particular site bacteria must adhere to host cells or to layers covering these cells (Ofek and Doyle, 1994f). The mucosal surfaces of the respiratory, gastrointestinal and urogenital tracts are the most common portals by which infectious bacteria enter the deeper tissues of a mammalian host. Thus, adhesion to the epithelial cells of these mucosal surfaces and then colonization of the mucosal tissue are considered the first stages on the infectious process. In numerous cases the adhesion is mediated by special protein molecules (known as adhesins) associated with proteinaceous organelles (known as fimbriae or pili). These adhesins, which are on the surface of the infectious bacteria, combine with complementary structures on the mucosal surfaces. Adhesion to mucosal surfaces offers the infectious agent a number of advantages. It allows the bacteria to firmly attach and thereby resist dislocation by the hydrokinetic forces that typically act on these surfaces. And it gives better access to nutrients as well as more protection from deleterious effects of antimicrobial agents in the surrounding milieu (Zafriri et al., 1987). Although adhesion is an important determinant of mucosal colonization, especially with respect to the animal and tissue tropism of the invading organism, several critical post-adhesion events are required for bacterial colonization. Triggered by the adhesion of the bacteria to their complementary receptors, these events include upregulation of virulence factor expression in the bacteria on the one hand and induction of physiological changes in the host cells on the other. Among the latter are cell proliferation, increased mucus secretion, endocytosis of adherent bacteria, and release of pro- and anti-inflammatory mediators by mucosal and sub-mucosal cells. In the present article, we review the current state of knowledge of bacterial adhesins and their mucosal cell receptors. We then discuss selected post-adhesion events and describe how they influence mucosal colonization and subsequent symptomatic infection. Finally, we show

how the knowledge gained provides a basis for the development of anti-adhesion agents that can block and even reverse bacterial colonization of mucosal surfaces before tissue damage.

Bacterial Adhesins and Their Cognate Receptors Types of Adhesin-Receptor Interactions The adhesive interactions of over 100 bacterial pathogens of humans and farm animals have been studied (Ofek and Doyle, 1994a; Karlsson, 1995). Based on these studies, three main types of adhesin-receptor interactions can be distinguished (Table 1). The first type, probably shared by the majority of bacterial pathogens is due to lectin-carbohydrate recognition. Many of the bacterial adhesins are lectins, a class of sugarbinding proteins that link the bacteria to carbohydrate moieties of glycoproteins or glycolipids on the mammalian host cell (Table 2). In some cases bacterial surface polysaccharides of either the capsule or the outermembrane lipopolysaccharides binds to cognate lectins on host cell’s (e.g., macrophage) surface (Ofek et al., 1995). The second type involves recognition of a protein on the bacteria by a complementary protein on the mucosal surface. The third type, and the one least well characterized, involves binding interactions between hydrophobic moieties of proteins on one cell with lipids ion the other cell, or between lipids on either cell (Ofek and Doyle, 1994b). Gents differ only in a single hydroxyl group, present on the acyl of the 4-NH group in the piglet-associated compound but absent in the pig-associated compound as well as in the glycolipids of humans. The subtle age-related change in the glycolipids may explain why E. coli K99 can cause diarrhea in piglets, but not in adult pigs or humans.

Multiple Adhesins A number of common themes have emerged regarding the interactions between bacteria and

CHAPTER 1.2

Bacterial Adhesion

17

Table 1. Molecular features of adhesin-receptor interactions in bacterial adhesion to host cells. Type of interaction

Bacterial ligand (and example)

Lectin-carbohydrate

Lectin (type 1 fimbriae)

Glycoprotein (uroplakin on bladder cells)

Wu et al., 1996

Polysaccharide (Klebsiella capsule)

Lectin (mannose receptor of macrophages)

Ofek et al., 1995

Protein-protein

Fibronectin binding proteins (F protein of S. pyogenes)

Fibronectin (fibronectin on respiratory cells)

Hanski et al., 1992; 1996

Hydrophobin-protein

Glycolipid (lipoteichoic acid of S. pyogenes)

Lipid receptors? (lipid-binding region of fibronectin on epithelial cells)

Courtney et al., 1990 Hasty et al., 1992

Lipid binding proteins (surface protein of Campylobacter spp.)

Membrane lipids (phospholipids and sphingolipids of cells)

Szymanski et al., 1996 Sylvester et al., 1996

mucosal cells. The most notable is the concept that pathogenic bacteria attach to mucosal cells typically through multiple adhesive interactions. Thus, a bacterial cell may express several adhesin moieties, each one specific for a distinct receptor molecule on the epithelial cell surface (for examples see Table 4). Interactions may be mediated by multiple bacterial adhesins that are structurally similar but may exhibit different binding specificities such as the type 1 and P fimbriae of uropathogenic E. coli (Table 3). Alternatively, adhesins may be structurally and chemically dis-

Receptor on host cell (and example)

References

similar, as is the case with the lipoteichoic acid (LTA) and proteinaceous adhesins of Streptococcus sanguis. Some pathogens (e.g., Neisseria gonorrhoeae) produce two surface lectins, each specific for distinct carbohydrate structures, one found in glycolipids and the other in glycoproteins. In many instances, different sub-populations of a bacterial clone express these distinct adhesins. By generating several phenotypic variants each expressing adhesins of distinct specificities, a given bacterial clone will increase the reper-

Table 2. Examples of carbohydrates as attachment sites for bacteria colonizing mucosal surfaces. Organism

Target tissue

E. coli type 1 P S CFA/1 CS3 K1 K99 H. pylori

Urinary tract Urinary tract Neural Intestinal Intestinal Endothelial Intestinal Stomach

N. gonorrkea

Genital

P. aeruginosa

Intestinal

H. influenza S. pneumoniae M. pneumoniae S. suis K. pneumoniae a

Respiratory Respiratory Respiratory Respiratory Respiratory Respiratory Respiratory & enterocytes

Carbohydrate structure

Formb

Mana3[Mara3(Mana6) Gala4Gal NeuAc(a2–3)Galb3GalNAc NeuAc(a2–8)GalNAcb4Gal GlcNAc(b1–4)GlcNAc NeuGc(a2–3)Gal(b1–4)Glc NeuAc(a2–3)Gal Lewis-b blood group Glucose-fatty acid Lactosyl ceramide Gal(b1–4)Glcb NeuAc(a2–3)Gal(b1–4)GlcNAc Gal(b1–3)GlcNAc, Fucose Mannose GalNAc(b1–4)Gal GalNAcb4Gal GalNAcb4Gal GlcNAcb3Gal NeuAc(a2–3)-Gal(b1–4)GlcNAc Gal(a1–4)Gal Gal(a1–4)Gal

Glycoprotein Glycolipid Glycolipid Glycoprotein Glycoproteinc Glycoprotein Glycolipid Glycolipid Glycoprotein Glycolipid Glycolipid Glycolipid Glycoprotein Glycoprotein Glycoprotein Glycoprotein Glycolipid Glycolipid Glycolipid Glycoprotein Glycoprotein Glycoprotein Glycoprotein

Based on Sharon and Lis, 1996; bOfek and Doyle, 1994 and Karlsson, 1995; cWenner et al., 1955.

18

I. Ofek, N. Sharon and S.N. Abraham

CHAPTER 1.2

Table 3. Types of receptor-adhesin relationship in bacterial adhesion to animal cells. Typea A.

B. C. D.

Receptor molecule

Animal cell

Adhesin molecule

Bacteria, (source)

Dr blood group antigen

Erythrocytes Erythrocytes Erythrocytes

Dra fimbriae AFA II F 1845 fimb

E. coli, (UTI) E. coli, (ETEC) E. coli, (pigs)

Fibronectin (NH2 terminal)

Epithelial cell

Lipoteichoic acid Fibronectin binding protein

S. pyogenes S. aureus

Glycolipid (Gala1–4Gal) Fibronectin

Uroepithelial cell

P fimbriae, FsoG P fimbriae, FsoF/H

E. coli, (pyelonephritis)

66 kDa Gp CD11/18 Gp CD 48Gpb Uroplakinc

Erythrocytes Neutrophils Macrophages Uroepithelial cell

Type 1 fimbriae Type 1 fimbriae Type 1 fimbriae Type 1 fimbriae

E. coli, (mannose sensitive)

Adapted from Ofek and Doyle, 1994. A. Target host cell express one receptor molecule that contain three attachment sites fro three different adhesins produced by three clones of bacteria; B. Two bacterial species express two distinct adhesins that bind the bacteria to the same receptor molecule on target host cell; C. The same bacterial clone produce a fimbrial structure comprised of two subunits, each bind the bacteria to distinct receptor on host target cell; D. The same adhesin bind the bacteria to similar attachment sites contained in different receptor molecules (isoreceptors) expressed by various host target cells. b Baorto et al., 1997; cWu et al., 1996. Gp, Glycoprotein. a

toire of its target tissues and perhaps also acquire antigenic variability that will enhance its ability to withstand the multifaceted defenses of the host (Ofek and Doyle, 1994b). This notion is exemplified by pyelonephritic isolates of E. coli which express either P fimbrial or type-1 fimbrial adhesin at any given time. Because transmission from one host to another is via the feacal-oral route, it was postulated that the pyelonephritogenic isolates may need the type 1 fimbriae mainly to transiently colonize the upper respiratory tract. Such colonies might then provide a constant source of bacteria entering the stomach and thus increase the chances for the incoming bacteria to colonize the intestine (Bloch et al., 1992). Once in the urinary tract, the bacteria seem to need the P fimbrial adhesins to adhere to the urinary tissues (Roberts et al., 1994; Win-

berg et al., 1995). In fact, the diverse types of fimbrial adhesins carried by various enterobacteria may determine by virtue of their distinct receptor specificity which of the unique niches along the intestine are colonized (Edwards and Puente, 1998). In those instances where multiple adhesins are expressed simultaneously on the same organism, each adhesin appears to complement the other functionally. For instance, the cell surface LTA and the M protein co-expressed on the surface of Streptococcus pyogenes have both been implicated in mediating bacterial binding to Hep-2 cells (Hasty et al., 1992; Courtney et al., 1997). Adhesion of S. pyogenes appears to involve a two-step process. The first step is mediated by the interaction of LTA with fibronectin molecules on the host cells (Hasty et al., 1992) and the second

Table 4. Selected bacterial clones expressing multiple adhesins. Bacterial clone

Source of isolation

E. coli

Pyelonephritis

S. saprophyticus

Urinary

N. gonorrhea

Urogenital

S. sanguis

Dental plaque hydrophobin

Adapted from Ofek and Doyle, 1994.

Adhesin

Characteristics

Type P Type 1 Gal-GlcNAc Lipoteichoic acid Pilus Opa protein Protein Fimbriae Protein Lipoteichoic acid

Fimbrial lectin Fimbrial lectin Peripheral lectin Fibrillar hydrophobin Pilin adhesin Outermembrane Peripheral Fimbrial adhesin Peripheral lectin Fibrillar hydrophobin

CHAPTER 1.2

by binding of the M protein to an as yet unidentified receptor on these cells (Courtney et al., 1997).

Adhesins as Lectins Table 2 presents a list of bacterial lectins, their molecular forms and their sugar specifities. Whenever known, their animal and organ specificities are also included. Methods are available for the detection and identification of sugar specificities (Goldhar, 1994, 1995; Sharon and Ofek, 1995). For further details the reader is referred to the review literature (Cassels and Wolf, 1995; Karlsson, 1995; Ofek and Doyle, 1994c). The lectin-mediated adhesion can be inhibited both in vitro and in vivo by either simple or complex carbohydrates that compete with the binding of the lectins to host-cell glycoproteins or glycolipids. In general the affinity of simple sugars (e.g., mono- or disaccharides) to the adhesins or lectins is low, in the millimolar range. Affinity can be increased several orders of magnitude by suitable chemical derivatization (Firon et al., 1987). Increase also can be obtained by attachment of the mono- or disaccharides to polymeric carriers, to form multivalent ligands (Lindhorst et al., 1997; Sharon, 1996; Sharon and Lis, 1997). Some bacterial lectins recognize not only terminal sugars but internal sequences as well. For example, the tip adhesin Pap G of P fimbriae recognizes internal Gala (1-4)-Gal sequences on cell surface glycolipids (Table 2). When the bacterial adhesin binds the pathogen to a cognate glycolipid, the ceramide group of the latter may contribute to the affinity of the interaction in some cases (e.g. Helicobacter pylori; Table 3). The study of bacterial lectins or adhesins, especially when these molecules are associated with fimbriae that are multi-subunit structures, has been hampered by difficulties in obtaining lectins in pure soluble form. Recently, however, a major breakthrough was achieved by preparing fusion proteins from the ZZ polypeptide of staphylococcal protein A and the amino terminal region of either PapG I, PapGII or PapGIII (Hansson et al., 1995). The three fusion proteins exhibited distinct fine sugar specificities identical with those of the parent fimbriae. It is anticipated that many of the fimbrial lectins will be purified and their combining sites identified using fusion to stabilize the proteins and preserve their carbohydrate-binding activity.

Bacterial Glycoconjugates as Adhesins Mammalian macrophages express lectins, which recognize complementary carbohydrate structures on bacterial surface and mediate nonopsonic phagocytosis of bacteria. Although phagocytosis, termed lectinophagocytosis, of a

Bacterial Adhesion

19

number of bacterial species was found to involve macrophage lectins, the surface glycoconjugates that mediate binding to the macrophage lectin have been identified for only a few bacteria. The mannose receptor of macrophages was found to recognize K. pneumoniae capsules that contain Mana2/3Man or Rhaa2/3Rha sequences and Mycobacterium tuberculosis that have arabinomannan on the surface (Athamna et al., 1991; Schlesinger et al., 1994). For comprehensive reviews on macrophage lectin and bacterial polysaccharide interaction in the infectious process, see Ofek et al. (1995), Ofek and Sharon (1988), Speert (1992, 1988); and Zwilling and Eisenstein (1994). It was suggested that lipo-oligosaccharide/ lipopolysaccharide (LOS/LPS) on the outer membrane of Gram-negative bacteria mediates adhesion to nonprofessional phagocytes (including mucosal cells) as well as to mucus constituents (Jacques, 1996; Nassif and Magdalene, 1995). The evidence for this effect is not conclusive and is based on the following observations: (1) epithelial cells bind less mutant strain (lacking the O side chain of LPS) than they do parental strains, and isolated LPS acts as inhibitor of the binding; (2) LPS isolated from Vibrio mimicus causes agglutination of rabbit erythrocytes (Alam et al., 1996); (3) the heptose-3-deoxy-Dmanno-2-octulosonic acid disaccharide present in the inner core of LPS is recognized by a lectinlike molecule on the plasma membrane of rat hepatocytes (Parent, 1990); and (4) the binding and internalization of Pseudomonas aeruginosa by corneal epihtelial cells requires intact innercore LPS with a terminal glucose residue (Zaidi et al., 1996). In a few cases interaction between a lectin on one bacterial cell and the lipooligosaccharide on another cell may mediate aggregation of the bacteria (Blake et al., 1995). The animal lectin galectin-3 was found to recognize bacterial lipopolysaccharides of Gramnegative bacteria (Mey et al., 1996). In no case, however, has there been definitive proof presented or identification made of a mucosal cell lectin that binds carbohydrates from pathogenic bacteria.

Adhesin-Receptor Relationship The adhesins of a number of bacterial pathogens and their cognate receptor on the host cells has been characterized in a considerable number of pathogenic organisms (reviewed in Ofek and Doyle, 1994b; Sharon and Lis, 1997). Several general features are notable (Table 2). One receptor may contain more than one attachment site that is specific for two or more adhesins. This is illustrated by the Dr blood group glycoprotein, which acts as receptor on host cell membrane for

20

I. Ofek, N. Sharon and S.N. Abraham

CHAPTER 1.2

three different clones of E. coli each one produces a distinct adhesin that binds to a different region of the Dr group molecule (Ofek and Doyle, 1994e). Another general feature is that two different pathogens, each expressing structurally distinct adhesins, can exhibit the same receptor specificity. This is the case with Staphylococcus aureus and S. pyogenes, both of which bind to the amino terminal region of fibronectin on mucosal cells. The adhesin on S. aureus is a fibronectin-binding protein, whereas that of S. pyogenes is lipoteichoic acid (Table 3). The finding that several different respiratory tract pathogens recognize the disaccharide GalNAcb4Gal is yet another example of the above (Table 2). It has been suggested that the GalNAcb4Gal sequence is preferentially accessible in glycolipids of the respiratory epithelium and this allows firm binding of a diverse group of respiratory pathogens bearing the suitable adhesins. In some cases, however, distinct adhesins share specificity but are carried by different bacteria that colonize different tissues and animal hosts, as is the case for the Gala (1-4)Gal-specific lectins of the uropathogenic Pfimbriated E. coli, the pig pathogen Streptococcus suis (Tikkanen et al., 1995), and the respiratory/ enteropathogenic P-like fimbriated K. pneumoniae (Prondo-Mordarska et al., 1996). Conversely, the same bacterial adhesin can bind to several distinct receptors on different cell types; such receptors are called isoreceptors. For instance several glycoproteins ranging in size from 110–45 kDa have been described as receptors for type 1 fimbriae on different cell types (Table 2). All these isoreceptor glycoproteins share a common oligomannose-containing attachment site for FimH, the adhesin subunit of type 1 fimbriae. Another situation is when an adhesin molecule contains multiple domains, each with distinct receptor specificity as is the case of the filamentous hemagglutinin adhesin of Bordetella pertussis. This hemagglutinin, which has been cloned and sequenced, contains at least three domains: (1) an arginine-glycine-aspartate (RGD)-containing sequence which binds the bacteria to a CR3 integrin present on pulmonary macrophages (Relman et al., 1989); (2) a carbohydrate-binding domains specific for galactose (Tuomannen et al., 1988) and (3) a carbohydratebinding domain specific for sulfated sugars (Menozzi et al., 1994).

glycosylated but to different extents. Included are structural glycoproteins that are typical constituents of the ECM such as collagens, elastin, fibronectin, fibrinogen, laminin, chondriotin sulfate proteoglycans and heparan sulfate proteoglycans. Many mucosal colonizers express adhesins that specifically recognize one or more of these substances. The same three categories of adhesin-receptor interactions, presented in Table 1, occur between bacteria and ECM components. They may be interactions between proteins only, between lipids and proteins, or between lectins and carbohydrates. A more thorough discussion of these ECM-bacteria interactions may be found in excellent reviews (Patti and Höök, 1994; Hasty et al., 1994; Patti et al., 1994; Wadstrom et al., 1994). Among the various ECM components, interactions with fibronectin have been studied the most at both the molecular and cellular levels. Because this multifunctional glycoprotein is found on the surface of many types of cells including mucosal ones, fibronectin probably acts as a receptor for bacterial adhesion and colonization. The adhesion of bacteria to extracellular matrix components other than fibronectin is becoming more appreciated. Examples of recent studies describing specific structures that mediate binding of bacteria to such betacomponents are shown in Table 5. A remarkable feature is that many of the bacterial species studied express on their surfaces at least two proteins that bind a specific ECM component. Thus, Helicobacter pylori expresses a laminin-specific adhesin that may be either a 25 kDa sialic-acidbinding lectin, which recognizes sialyl residues of laminin, or a lipopolysaccharide which recognizes other, as yet unidentified, regions in laminin (Valkonen et al., 1994, 1997). Many studies have established fibronectin as an important receptor for S. pyogenes and other bacteria on mucosal surfaces (Ofek and Doyle, 1994e; Courtney et al., 1990). At least six different molecules on S. pyogenes surfaces were found to recognize fibronectin, including LTA, protein F/Sfb, a 28 kDa fibronectin-binding protein, glyceraldehyde-3-phosphate dehydrogenase, serum opacity factor and a 54 kDa fibronectin-binding protein (FBP54; reviewed in Hasty and Courtney, 1996). It is not clear whether all these fibronectin-binding entities mediate the adhesion of streptococci to mucosal surfaces.

Interaction of Bacterial Adhesins with Extracellular Matrix

Consequences of Bacterial Adhesion to Cells and Tissues

Mucosal cells are often covered by a layer referred to as extracellular matrix (ECM), which is a heterogeneous assembly of proteins, mainly

Recently it has been shown that adhesins not only enable colonization of mucosal surfaces but also elicits a variety of distinct responses in the

CHAPTER 1.2

Bacterial Adhesion

21

Table 5. Examples of bacterial adhesins mediating binding of the bacteria to ECM glycoproteins. Bacteria Borrelia burgdorferi H. influenzae N. gonorrhea P. aeruginosa Staphylococcus aureus

Mycobacterium bovis E. coli Bordetella pertussis H. pylori Listeria monocytogenes

Bacterial adhesin

ECM component

References

19 and 20kDa proteins Protein A (Osp A) and 70kDa protein P2 and P5 outermembrane proteins Opa protein 57 and 59kDa outermembrane proteins 42–48 and 77–85kDa outermembrane proteins and Flagellar 65.9kDa FLi F (MS ring) 138 and 127 surface proteins Cna protein (55Kda domain) ClfA (clumping factor) FnBPA and FnBPB 28kDa protein Gaf D protein of G fimbriae Filamentous hemagglutinin (N-terminal region of FHA) Lipopolysaccharide and 25kDa protein ActA outermembrane protein

Proteoglycan decorin Plasminogen Respiratory mucin Proteoglycan Laminin Respiratory mucins

1, 1a 2 3, 3a, 3b 4 5 6, 6a

Nasal mucin Collagen Fibrinogen Fibronectin Heparan Laminin Heparan Laminin Heparan

7

8 9 10 11, 11a 12

Key to references: 1. Guo et al., 1995; 1a. Leong et al., 1995; 2. Hu et al., 1995; 3. Davis et al., 1995; 3a. Reddy et al., 1996; 3b. Kubiet and Ramphal, 1995; 4. Putten and Paul, 1995; 5. Plotkowski et al., 1996; 6. Scarfnman et al., 1996; 6a. Akora et al., 1996; 7. Shuter et al., 1996; Foster and Hook, 1998; 8. Menozzi et al., 1996; 9. Saarela et al., 1996; 10. Hannah et al., 1994; 11. Valkonen et al., 1994; 11a. Valkonen et al., 1997; 12. Alvarez-Domínguez et al., 1997.

host cells as well as in the bacteria which can markedly affect the course of the infectious process (reviewed in Finlay and Cossart, 1997). In this section, selected examples are presented to illustrate the above notion.

Induction of Bacterial Virulence Genes The urinary tract is relatively refractory to bacterial colonization. In addition to resisting the constant hydrokinetic forces acting in this organ, a potential pathogen must multiply fast enough in urine to compensate for the diluting effects of the latter. Urine is a complex fluid containing a variety of excreted products but is growth limiting for bacteria, in part, because it is low in free iron. The intrinsic iron acquisition machinery of uropathogenic E. coli is activated upon complex formation between the PapG fimbrial adhesin with its Gala (1-4)Gal-containing globoseries receptor (Zhang and Normark, 1996). When P-fimbriated bacteria attached to immobilized receptor, transcriptional activation of a sensorregulator protein, AirS, was detected. This sensor protein, located in the cytoplasmic membrane, belongs to the two-component family of signal transduction factors. The precise mechanism of AirS action is as yet not known. It is believed to regulate the bacterial iron acquisition system and iron-regulated membrane proteins to facilitate the translocation of iron into the bacterium. Uropathogenic E. coli, in which the airS gene was

knocked out, lost its capacity to grow in urine. It would appear that uropathogenic bacteria can “sense” receptors (e.g. of the globoseries) in the urinary tract environment via PapG and respond by colonizing this body site. These findings point to an intriguing new function for bacterial P fimbriae, namely, that of a sensory organelle. The strategic location of PapG at the distal tips of the peritrichously arranged fimbriae probably facilitates this purported role. This finding is one of an increasing number of cases showing that bacterial pathogens are intrinsically capable of responding to cues from host cells following interactions between complementary cellsurface molecules (Cotter and Miller, 1996; Finlay and Cossart, 1997). In addition, these observations provide a molecular basis for earlier findings. Various bacteria obtain a growth advantage after attachment to host cells, as demonstrated for type 1 fimbriated E. coli and N. gonorrhoeae, which exhibit shorter lag periods when adhering to tissue culture cells (Zafriri et al., 1987; Bessen and Gotschlich, 1986).

Induction of Cytokine Release from Mucosal Cells In addition to evoking responses in the adherent bacteria, the specific coupling of the bacterial adhesins with their receptors also elicits a range of mucosal cell responses (Bliska and Falkow, 1992). For example, adhesion of the P-fimbrial adhesin to its receptors on mouse uroepithelial cells elicits the release from these cells of several immunoregulatory cytokines including inter-

22

I. Ofek, N. Sharon and S.N. Abraham

leukins (ILs)-1a-, b, -6 and -8 (Svanborg et al., 1996). It also triggers intracellular release of ceramides that may be derived from the globoseries receptor itself or from neighboring sphingomyelin molecules by the action of endogenous sphingomyelinases (Hedlund et al., 1996; Svanborg et al., 1996). Ceramide is known to be a critical second messenger in signal transduction processes capable of activating the Ser/Thr family of protein kinases and phosphatases and leading eventually to cytokine production. This bacterial adhesin- mediated mechanism of signaling is reminiscent of that utilized by immunoregulatory cytokines such as tumor necrosis factor alpha (TNFa) and IL-1 when evoking cellular responses (Svanborg et al., 1996). Thus bacterial adhesin appears to be functionally mimicking the host’s immunoregulatory molecules. Although the type 1 fimbriae of uropathogenic E. coli also stimulate a cytokine response from uroepithelial cells, the array of cytokines released is different from those elicited by P fimbriae (Connell et al., 1996b). The transmembrane signaling pathway of cytokine release by type 1 fimbriae has not been investigated but its clarification could benefit from the recent identification of uroplakin as the putative FimHreceptor on epithelial cells (Wu et al., 1996). Adhesion of Gram-positive bacteria to epithelial cells may also cause release of cytokines from the cells. For instance, group A streptococci adherent to HEp-2 cells via both M protein and LTA adhesins cause release of IL-6 from the target cells (Courtney et al., 1997). Perhaps more interesting are the findings that interaction of bacteria with ECM constituents may also trigger signal transduction in the underlying host cells (Juliano and Haskill, 1993).

Induction of Cytokine Responses in Inflammatory Cells The capacity of bacterial adhesins to elicit cytokine responses is not confined to mucosal cells. Lectinophagocytosis mediated by fimbriae such as type 1 fimbriae of E. coli or of type 2 fimbriae of Actinomyces viscosus is associated with stimulation of the phagocytic cells (Sandberg et al., 1988; Ofek et al., 1995). Indeed, type 1 fimbriae of uropathogenic E. coli are capable of binding to and eliciting immunoregulatory products from a wide range of inflammatory cells including macrophages, neutrophils, mast cells, and B and T lymphocytes in vitro (reviewed in Connell et al., 1996a). That these interactions may occur in vivo with significant physiologic effects is suggested by experiments in which mice injected intraperitoneally with type 1 fimbriated E. coli generated lysosomal b-N-acetylglucosaminidase and a large spike of TNFa in the peritoneal fluid

CHAPTER 1.2

(Bernhard et al., 1992; Malaviya et al., 1996). The fimbrial adhesin, FimH, plays a key role in this exposure because intraperitoneal challenge with a FimH-minus isogenic mutant resulted in only a limited TNFa response (Malaviya et al., 1996). The source of TNFa in the mouse peritoneum was determined to be mast cells because mice genetically deficient in these cells exhibited a limited TNFa response following intraperitoneal injection of type 1 fimbriae. Notably, this TNFa response was accompanied by a large influx of neutrophils into the peritoneum, consistent with the fact that TNFa is a potent neutrophil chemoattractant (Malaviya et al., 1996). Thus, one of the immediate outcomes of type 1 fimbriae-mediated activation of mast cells is recruitment of neutrophils to sites of bacterial challenge. Because mast cells are found preferentially in mucosal surfaces, the interaction of type 1 fimbriae of E. coli with such cells could contribute to the influx of neutrophils from surrounding blood vessels leading to the translocation of the bacteria through the epithelial barrier and subsequent entry into the lumen. The excessive transepithelial migration of neutrophils during infections may predispose this barrier to increased bacterial penetration (Finlay and Cossart, 1997) and raises the possibility that facets of the host’s immune response may be co-opted by pathogenic bacteria to enhance their virulence.

Impact of Bacteria-Elicited Inflammatory Responses Evaluating the physiologic effects of some of the adhesin-elicited cytokines at sites of bacterial infection is difficult because these effects are numerous and complex (Abraham and Malaviya, 1997; Henderson et al., 1996). Some of the responses evoked in the mucosa following the adherence of pathogenic bacteria include increased mucus secretion, proliferation of epithelial cells and recruitment and activation of a variety of phagocytic cells. All of these responses could potentially affect the early elimination of the pathogen (Abraham and Malaviya et al., 1997; Henderson et al., 1996). However, some of the adhesin-triggered secreted products of host cells may have severe pathophysiologic effects on the surrounding tissue, particularly when released in excess or at inopportune times (Abraham and Malaviya, 1997). Although direct evidence is still lacking, considerable circumstantial evidence supports the notion that the many proteases, oxygen radicals, and cytotoxic cytokines secreted after inflamatory cells are activated by type 1 fimbriated E. coli (Tewari et al., 1994; Malaviya et al., 1994, 1996) are detrimental to the host and foster bacterial pathogenesis. For example, the elastases, oxygen radicals and other

CHAPTER 1.2

cytotoxic agents, released from neutrophils following their interaction with type 1 fimbriae of E. coli in the kidney, are major contributors to renal scarring (Steadman et al., 1988; Topley et al., 1989). Whether an inflammatory response favors the host or pathogen may depend on other prevailing factors including the host’s immune status and the intrinsic virulent capabilities of the pathogen. The number of bacteria at the site of infection may be another critical factor in light of the recent findings that certain bacteria have “quorum sensing” ability (Passador et al., 1993; i.e., they sense their population density at a given site and, upon reaching a critical density, coordinately turn on the expression of a battery of new virulence factors.)

Bacterial Uptake by Phagocytes In addition to inducing the release of pharmacologically active mediators from various host cells, bacterial adhesins also elicit the phagocytic uptake of bacteria under serum-free conditions (reviewed in Ofek et al., 1995). The process involves a number of molecular mechanisms; as mentioned this process has been termed lectinophagocytosis, in analogy to opsonophagocytosis (Ofek and Sharon, 1988; Ofek et al., 1995). The best-characterized system of lectinophagocytosis is that of bacteria carrying the mannosespecific type 1 fimbrial lectins. The fact that a bacterial adhesin that promotes bacterial colonization and infection may also promote ingestion by phagocytic cells would seem a paradox. Although earlier work showed that bacteria are occasionally killed by the phagocytes, new evidence has emerged to suggest that type 1 fimbriae-elicited bacterial phagocytosis by macrophages may actually benefit the bacterial population (Baorto et al., 1997). In vitro survival assays in macrophages revealed that, unlike E. coli phagocytized via opsonin-mediated processes, E. coli phagocytized via type 1 fimbriae survived much of the intracellular killing. It has been suggested that by associating with CD48, a glycosylphosphoinositol-linked moiety on the surface of macrophages, the bacteria gain access to a lipid processing pathway that bypasses the normal phagocytic killing mechanisms of the macrophages (Baorto et al., 1997). This finding provides a molecular basis for earlier observations showing that, compared to bacteria ingested via opsonophagocytosis, bacteria subjected to lectinophagocytosis are often markedly less sensitive to killing by phagocytes (reviewed in Ofek et al., 1995). It is noteworthy that lectinophagocytosis comes into play only at body sites where opsonizing is poor such as in the urinary mucosa.

Bacterial Adhesion

23

Internalization by Nonphagocytic Cells Contact between bacterial adhesins and complementary receptors on so called nonphagocytic cells can trigger internalization of adherent bacteria (reviewed by Finlay and Falkow, 1990, 1997; Marra and Isberg, 1996). This has been demonstrated with such classical intracellular pathogenic species as Listeria, Yersinia, Shigella, Salmonella and Bartonella (Table 7). These organisms enter and proliferate in nonphagocytic cells in vitro and in vivo. Probably because of the development of highly sensitive and reproducible techniques to measure bacterial entry into mammalian cells (Tang et al., 1993), several well-known “extracellular” pathogens have recently been reported to be capable of penetrating nonphagocytic cells (e.g., epithelial and endothelial cells) and of surviving for a limited period and, in some cases, even of proliferating intracellularly. Unlike the classical or professional intracellular pathogens, entry of the extracellular pathogens is usually limited to a subset of bacterial strains within the same species, probably because entry into requires the co-expresion of multiple components such as adhesins and constituents of the secretory system (De Vries et al., 1996). Furthermore, the capacity to enter nonphagocytic cells is not necessarily associated with virulence of the extracellular pathogen. For example, isolates from carrier-state or nonencapsulated strains of S. pyogenes can invade epithelial cells, whereas pharyngitis isolates (Sela, 1998) or virulent encapsulated strains (Schrager et al., 1996) invade poorly. Excluding the classical intracellular pathogens, the list of bacterial species capable of invading nonphagocytic cells includes Actinobacillus actinomycetemcomitans (Meyer et al., 1996), Pseudomonas aeruginosa (Fleiszig et al., 1995, 1996), Burkholderia (Pseudomonas) cepacia (Burns et al., 1996), E. coli (Meier et al., 1996; Jouve et al., 1997; Donnenberg et al., 1997; Goluszko et al., 1997), K. pneumoniae (Oelschlaeger and Tall, 1997), N. gonorrhoeae (Weel et al., 1991), N. meningitidis (Virji et al., 1993), Porphyromonas gingivalis (Weinberg et al., 1997), Streptococcus agalactiae (Hulse et al., 1993; Valentin-Weigand et al., 1997; Gibson et al., 1993), S. aureus (Vann et al., 1987; Hamill et al., 1986) and S. pyogenes (Greco et al., 1995; LaPenta et al., 1994). Conceivably, the ability to enter nonphagocytic cells is an integral part of the pathogenic process of many infectious bacteria. While the classical intracellular pathogens utilize this ability to spread from cell to cell and to penetrate into deep tissue, other pathogens may utilize this trait to temporarily hide from the host’s immune cells or from antibiotics. Thus, bacteria surviving within nonphagocytic cells

24

I. Ofek, N. Sharon and S.N. Abraham

CHAPTER 1.2

Table 6. Inhibitors of bacterial lectin/adhesin as anti-adhesion drug for preventing infection in experimental animals. Inhibitor

Bacteria

Mannose or its glycosides

E. coli type 1

Gala4Galb containing oligosaccharide

K pneumoniae type 1 Shigella flexneri type 1 E. coli type P

Glycopeptides (from serum glycoproteins) Galactose, mannose and N-acetylglucosamine Sialyl containing oligosaccharide GalNAcb4Gal containing oligosaccharide N-Acetylglucosamine

E. coli K99 P. aeruginosa H. pylori S. pneumoniae S. pneumoniae

Animal

Site of infection

Mice Mice Rats Guinea pigs Mice Monkeys Calves Human Piglet Rabbit Mouse

Bladder Gut Bladder Eye Urinary tract Urinary tract Gut Ear Gut Lung Lung

Gal, galactose; GalNAc, N-acetylgalactosamine. Adapted from Sharon, 1996.

a

serve as a critical reservoir from which reinfection of the host can take place. The mechanisms employed by various bacteria to gain access into nonphagocytic cells are diverse and often complex (see range of molecules implicated in bacterial invasion of host cells in Table 7). For example, Yersinia enterolitica

employs a single cell surface protein, invasin, whose cognate receptors on the host cell membrane are b integrins (Isberg, 1996). When invasin binds with high affinity to b integrins, the close association between the integrins and cytoskeletal elements of the cell membrane triggers the bacterial uptake. Particles that are

Table 7. Examples of bacteria capable of invading nonphagocytic cells. Surface constituents for Bacteria (reference)

adhesion

Listeria monocytogenes (1)

?

Yersinia spc (2) Salmonella sp (3) Shigella sp (4) Bartonella sp. (5)

Ail protein YadA protein ? ? BFP

EPEC, STECd (6) E. coli (7) N. gonorrhea (8)

BFPe AfaIII Pili

S. pyogenes (9)

LTA

K. pneumoniae (10)

Type 1 fimb.

entrya

Receptor for entry

Intracellular proliferation

Tissue damageb

InlA InlB ActA Invasinb

E cadherin

+

+

Proteoglycan Integrins

+

+

Sip proteins Ipab proteins IalA IalB Intamin AfaE, AfaD proteins Opa A,C

CD42 Integrins Glycolpid

+ + +

+ + +?

Integrin, HP90 ? CD66 family Vitronectin Heparan sulfate Fibronectin

-? -

+ ? +

-

-

GlcNAcf

NT

-

F protein M protein ?

Key to references (1) Gaillard et al., 1991; Mengaud et al., 1996; (2) Iseberg et al., 1987; Iseberg and Leong, 1990; Miller and Falkow, 1998; Saltman et al., 1996; Schulze-Koops et al., 1992; 1993; (3) Chen et al., 1996; Francis et al., 1993; (4) Watarai et al., 1995, Mennardi et al., 1996; Zychlinski and Sansonetti, 1997; (5) Minnick et al., 1996; (6) Donnenberg et al., 1992, 1997; Frankel et al., 1995, 1996; Rosenshine et al., 1996; Paton and Paton, 1998; (7) Jouve et al., 1997; (8) Weel et al., 1991, Makino et al., 1991, van Putten et al., 1995, Virji et al., 1996; Chen et al., 1997; Gomez-Durate et al., 1997; (9) LaPenta et al., 1994; Greco et al., 1995; Jadoun et al., 1997; Molinary et al., 1997; (10) Oelschlaeger and Tall, 1997; Fumagalli et al., 1997. a The surface constituents required for entry usually can function as adhesins as well. b Damage usually associated with inflammation resulting from the entry process (Shigella) or from direct damage of the cell membrane of the target host cells (e.g. E. coli and Salmonella). c Enteropathogenic Yersinia species e.g. Y. enterolytica and Y. pseudotuberculosis. d Enteropathogenic E. coli. Entry was documented only in tissue cell culture (Donnenber et al., 1990), but the intimin is required for intimate association and induction of the effacement/attaching lesion. e Bundle forming pili. f N-Acetylglucosamine containing glycoprotein on tissue culture cells.

CHAPTER 1.2

coated with invasin proteins (or functionally relevant portions of the protein) and exposed to tissue culture cells are readily internalized by the cells. More complex modes of entry requiring specific secretion systems of the bacteria have been reported for certain enteropathogenic E. coli (Javris et al., 1995) and species of Shigella (Allaoui et al., 1993), Salmonella (Ginocchio et al., 1992) and Bartonella (Minnick et al., 1996). Perhaps the most remarkable of these systems involves enteropathogenic E. coli (EPEC) where the type III secretory system of the bacteria inserts into the host cell membrane a protein (HP90) that serves in turn as the receptor for the bacterial adhesin (Kenny et al., 1997; Nataro and Kaper, 1998). The process of internalization involve sequential interactions between EPEC and the host cell. The first step of adhesion occurs via bundle fimbria and is followed by intimate contact via a second adhesin termed intimin. The receptor on the host cell for the intimin is HP90, which is produced by the bacteria, phosphorylated, and then inserted into the host cell membrane by the type III secretory system of the bacteria. With the binding of intimin, the bacteria become internalized by the host cells. A similar mechanism was described for the internalization of pathogenic Neisseria by nonphagocytic cells (Dehio et al., 1998). In another recently reported mechanism, the bacteria after adhesion to their cognate receptor initiate a signaling cascade resulting in activation of phosphatidylcholine-specific phospholipase C and acidic sphingomyelinase, to allow entry of N. gonorrhoeae into nonphagocytic cells (Grassme et al., 1997). Finally, in some cases the molecular mechanism utilized by the bacteria to gain entry into nonphagocytic cells appears to be the same as that involved in the uptake of bacteria by phagocytes. A case in point is the specific interaction between CD66 on the mammalian cell surface and the N. gonorrhoeae Opa proteins that triggers the uptake of bacteria by both epithelial cells and polymorphonuclear cells (GrayOwen et al., 1997; Virji et al., 1996; Chen et al., 1997; Sauter et al., 1993).

Concluding Remarks Experiments in animals have proven that it is possible to prevent infections by blocking the adhesion of the pathogen to target tissue. These findings have stimulated the development of antiadhesion drugs for preventing and treating microbial infections in humans (reviewed in Kahane and Ofek, 1996). New classes of these drugs are greatly needed because of the increasing incidence of pathogenic organisms resistant to conventional antibiotics. It is believed that strains

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with genotypic resistance to the anti-adhesion agents will spread much slower than strains resistant to conventional drugs, such as antibiotics aimed at killing the organisms. The reason is that both anti-adhesion-sensitive and -resistant strains are shed to continue transmission from host to host, whereas only antibiotic-resistant strains are transmitted following therapy. Because lectin-mediated adhesion is a mechanism shared by many pathogens most investigators have focused their efforts to prevent bacterial infections on blocking the pathogen’s lectins. The preferable target site is the mucosal surfaces where phagocytic cells are scarce and where most infections are initiated. A number of strategies have been suggested including enhancement of mucosal immunity by s-IgA anti-adhesin antibody induction, use of metabolic inhibitors of adhesin expression (e.g. sublethal concentration of antibiotics), and of dietary inhibitors, in particular receptor analogs (reviewed in Ofek and Doyle, 1994b; Kahane and Ofek, 1996). In the latter strategy, the lectin or adhesin is inhibited by sugars for which the lectin is specific (Table 6). This was first demonstrated in the late 1970s, when it was shown that methyl a-mannoside can protect mice against urinary tract infection by type 1 fimbriated E. coli; methyl a-glycoside which is not recognized by the bacteria, was not effective (Aronson et al., 1979). Subsequent studies by many other groups have proven beyond any doubt the drug potential of anti-adhesive compounds (Table 6; Beuth et al., 1995; Sharon, 1996; Ofek and Sharon, 1990; Zopf and Roth, 1996). Thus, derivatives of galabiose that inhibit the adhesion of P fimbriated E. coli to animal cells in vitro, prevented bacterial infections in the urinary tract of mice and monkeys. Antibodies against mannosecontaining compounds present on epithelial cells prevented urinary tract infection in mice by type 1 fimbriated E. coli. in mice, and orally administered sialylated glycoproteins protected colostrum-deprived newborn calves against lethal doses of enterotoxigenic E. coli K99. In a clinical trial in humans, patients with otitis externa (a painful swelling with secretion from the external auditory canal) caused by P. aeruginosa were treated with a solution of galactose, mannose and N-acetylneuraminic acid (Beuth et al., 1996). The results were fully comparable to those obtained with conventional antibiotic treatment. An attractive candidate is oligosaccharides such as those found in human milk and other body fluids, that have been shown to inhibit the adhesion to cells and tissues of strains of H. pylori and S. pneumoniae (Zopf et al., 1996; Simon et al., 1997). Human milk is a potential source of inhibitors of bacterial adhesion because it is rich in disac-

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charides that may act as receptor analogs (Ashkenazi, 1996). However, other dietary constituents also may exhibit anti-adhesion activity and may be used to prevent bacterial infections. For example, cranberry juice contains at least two inhibitors of uropathogenic E. coli (Ofek et al., 1991) and according to one well documented report, it reduced the incidence of urinary tract infections in elderly women (Avorn, et al., 1994). These findings illustrate the great potential of inhibitors of adhesion in the prevention and perhaps also treatment of bacterial infections. Moreover, they raise hopes for the development of anti-adhesive drugs for human use. The development of anti-adhesion therapy targeted at the microbial lectins has been hampered by the great difficulty in large-scale synthesis of the required inhibitory saccharides. An alternative is glycomimetics, compounds that structurally mimic the inhibitory carbohydrates, but which may be more readily obtainable. Eventually, a cocktail of inhibitors, or a polyvalent one, will have to be used, since many infectious agents express multiple specificities. The design of such drugs will certainly benefit from more detailed information about the specificity of the microbial surface lectins and the elucidation of the atomic structure of their combining sites, none of which is yet known.

Literature Cited Abraham, S. N., and Malaviya, R. 1997. Mast cells in infection and immunity. Infect. Immun. 65:3501–3508. Abraham, S. N., Sun, D., Dale, J. B., and Beachey, E. H. 1988. Conservation of the mannose-adhesin protein among type 1 fimbriated members of the family Enterobacteriaceae. Nature 336:682–684. Akora, S. K., Ritchings, B. W., Almira, E. C., Lory, S., and Ramphal, R. 1996. Cloning and characterization of Pseudomonas aeruginosa FliF, necessary for flagellar assembly and bacterial adherence to mucin. Infect. Immun. 64:2130–2136. Alam, M., Miyoshi, S-I., Tomochika, K-I., and Shinoda, S. 1996. Purification and characterization of novel hemagglutinins from Vibrio mimicus: a 39-kilodalton major outer membrane protein and lipopolysaccharide. Infect. Immun. 64:4035–4041. Allaoui, A., Sansonetti, P. J., and Parsot, C. 1993. MxiD, an outer membrane protein necessary for the secretion of the Shigella flexneri Ipa invasins. Mol. Microbiol. 7:59– 68. Alvarez-Domínguez, C., Vázquez-Boland, J-A., CarrascoMarín, E., López-Mato, P., and Lyva-Cobian, F. 1997. Host cell heparan sulfate proteoglycans mediated attachment and entry of Listeria monocytogenes, and the listerial surface protein ActA is involved in heparan sulfate receptor recognition. Infect. Immun. 65:78–88. Aronson, M., Medalia, O., Schori, L., et al. 1979. Prevention of colonization of the urinary tract of mice with Escher-

CHAPTER 1.2 ichia coli by blocking of bacterial adherence with methyl a-D-mannopyranoside. J. Infect. Dis. 139:329–332. Athamna, A., Ofek, I., Keisari, Y., Markowitz, S., Dutton, G. G. S., and Sharon, N. 1991. Lectinophagocytosis of encapsulated Klebsiella pneumoniae mediated by surface lectins of guinea pig alveolar macrophages and human monocyte-derived macrophages. Infect. Immun. 59:1673–1682. Avorn, J., Monane, M., Gruwitz, J. H., Glynn, R. J., Choodnovskiy, I., and Lipsitz, A. 1994. Reduction of bacteriuria and pyuria after ingestion of cranberry juice. J. Amer. Med. Assoc. 271:751–754. Baorto, D. M., Gao, Z., Malaviya, R., Dustin, M., Van der Merwe, A., Lublin, D., and Abraham, S. N. 1997. Survival of FimH-expressing enterobacteria in macrophages relies on glycolipid. Nature 283:636–639. Baorto, D. M., Gao, Z., Malaviya, R., Dustin, M., Van der Merwe, A., Lublin, D., and Abraham, S. N. 1997. Survival of FimH-expressing enterobacteria in macrophages relies on glycolipid. Nature 283:636–639. Bernhard, W., Gbarah, A., and Sharon, N. 1992. Lectinophagocytosis of type 1 fimbriated (mannose-specific) Escherichia coli in the mouse peritoneum. J. Leukocyte Biol. 93:1645–1653. Bessen, D., and Gotschlich, E. C. 1986. Interaction of gonococci with Hela cells: attachment, detachment, replication, penetration and the role of protein II. Infect. Immun. 54:154–160. Beuth, J., Ko, H. L., Pulverer, G., Uhlenbruck, G., and Pichlmaier, H. 1995. Importance of lectins for the prevention of bacterial infections and cancer metastasis. Glycoconjugate J. 12:1–6. Kahane, I., and Ofek, I., Editors 1996. Toward anti-adhesion therapy of microbial diseases. Plenum Publishing Company. New York, NY. 408:297. Blake, M. S., Blake, C. M., Apicella, M. A., and Mandrell, R. E. 1995. Gonococcal opacity: lectin-like interactions between Opa proteins and lipopolysaccharide. Infect. Immun. 63:1434–1439. Bliska, J. B., and Falkow, S. 1992. Signal transduction in the mammalian cell during bacterial attachment and entry. Cell 73:903–920. Bloch, C. A., Stocker, A. D., and Orndorff, P. E. 1992. A key role for type 1 pili in enterobacterial communicability. Mol. Microbiol. 6:697–701. Burns, J. L., Jonas, M., Chi, E. Y., Clark, D. K., Berger, A., and Griffith, A. 1996. Invasion of respiratory epithelial cells by Burkholderi (Pseudomonas) cepacia. Infect. Immun. 64:4054–4059. Cassels, F. J., and Wolf, M. K. 1995. Colonization factors of diarrheagenic E. coli and their intestinal receptors. J. Industrial Microbiol. 15:214–226. Chen, T., Grunert, F., Medina-Marino, A., and Gotschlich, E. C. 1997. Several carcinoembryonic antigens (CD66) serve as receptors for gonococcal opacity proteins. J. Exp. Med. 185:1557–1564. Chen, L. M., Hobbie, S., and Galan, J. E. 1996. Requirement of CD42 for Salmonella-induced cytoskeletal and nuclear response. Science 274:2115–2118. Connell, I., Agace, W., Klemm, P., Schembri, M., Marild, S., and Svanborg, C. 1996a. Type 1 fimbrial expression enhances Escherichia coli virulence for the urinary tract. Proc. Natl. Acad. Sci. (USA) 93:9827–9832. Connell, I., Agace, W., Hedlund, M., Klemm, P., Schembri, M., and Svanborg, C. 1996b. Fimbriae-mediated adherence induces mucosal inflammation and bacteriurial

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Prokaryotes (2006) 2:32–85 DOI: 10.1007/0-387-30742-7_3

CHAPTER 1.3 ehT

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The Phototrophic Way of Life JÖRG OVERMANN AND FERRAN GARCIA-PICHEL

Introduction Photosynthesis is the utilization of radiant energy for the synthesis of complex organic molecules. The phototrophic way of life implies the capture of electromagnetic energy (see Light Absorption and Light Energy Transfer in Prokaryotes in this Chapter), its conversion into chemical energy (see Conversion of Light into Chemical Energy in this Chapter), and its use for cellular maintenance and growth (see Efficiency of Growth and Maintenance Energy Requirements in this Chapter). Photosynthesis may encompass the reduction of carbon dioxide into organic molecules, a mode of growth defined as photoautotrophy. The solar electromagnetic energy reaching the Earth’s surface (160 W·m–2; see Light energy and the spectral distribution of radiation) surpasses the energy contributed by all other sources by four to five orders of magnitude (electric discharge, radioactivity, volcanism, or meteoritic impacts; ~0.0062 W·m–2 on primordial Earth; Mauzerall, 1992; present day geothermal energy ~0.0292 W·m-2; K. Nealson, personal communication). At present the flux of electromagnetic energy supports a total primary production of 172.5 ¥ 109 tons dry weight·year-1 (168 g C·m-2·year-1; Whittaker and Likens, 1975). If this global primary production is converted to energy units (39.9 kJ·g C-1, assuming that all photosynthetic products are carbohydrate), 0.21 W·m-2 and thus 0.13% of the available solar energy flux are converted into chemical energy. Even at this low efficiency, the chemical energy stored in organic carbon still exceeds geothermal energy by at least one order of magnitude. As a consequence, photosynthesis directly or indirectly drives the biogeochemical cycles in all extant ecosystems of the planet. Even hydrothermal vent communities, which use inorganic electron donors of geothermal origin and assimilate CO2 by chemolithoautotrophy (rather than photoautotrophy), still depend on the molecular O2 generated by oxygenic phototrophs outside of these systems (Jannasch, 1989). Several lines of evidence indicate that in the early stages of biosphere evolution, prokaryotic

organisms were once responsible for the entire global photosynthetic carbon fixation. Today, terrestrial higher plants account for the vast majority of photosynthetic biomass; the chlorophyll bound in light-harvesting complex LHCII of green chloroplasts alone represents 50% of the total chlorophyll on Earth (Sidler, 1994). In contrast, the biomass of marine primary producers is very low (0.2% of the global value). However, the biomass turnover of marine photosynthetic microorganisms is some 700 times faster than that of terrestrial higher plants. Thus, marine photosynthetic organisms contribute significantly to total primary productivity (55·109 tons dry weight·year-1, or 44% of the global primary production). Because the biomass of cyanobacterial picoplankton (see Habitats of Phototrophic Prokaryotes in this Chapter) can amount to 67% of the oceanic plankton, and their photosynthesis up to 80% in the marine environment (Campbell et al., 1994; Goericke and Welschmeyer, 1993; Liu et al., 1997; Waterbury et al., 1986), prokaryotic primary production is still significant on a global scale. A single monophyletic group of marine unicellular cyanobacterial strains encompassing the genera Prochloroccoccus and Synechococcus with a global biomass in the order of a billion of metric tons (Garcia-Pichel, 1999) may be responsible for the fixation of as much as 10–25% of the global primary productivity. Additionally, prokaryotic (cyanobacterial) photosynthesis is still locally very important in other habitats such as cold (Friedmann, 1976) and hot deserts (Garcia-Pichel and Belnap, 1996) a nd hypertrophic lakes. Today, the significance of anoxygenic photosynthesis for global carbon fixation is limited for two reasons. On the one hand, phototrophic sulfur bacteria (the dominant anoxygenic phototrophs in natural ecosystems) form dense accumulations only in certain lacustrine environments and in intertidal sandflats. The fraction of lakes and intertidal saltmarshes which harbor anoxygenic phototrophic bacteria is unknown, but these ecosystems altogether contribute only 4% to global primary production (Whittaker and Likens, 1975). In those lakes harboring pho-

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totrophic sulfur bacteria, an average of 28.7% of the primary production is anoxygenic (Overmann, 1997). Consequently, the amount of CO2 fixed by anoxygenic photosynthesis must contribute much less than 1% to global primary production. On the other hand, anoxygenic photosynthesis depends on reduced inorganic sulfur compounds which originate from the anaerobic degradation of or ganic carbon. Since this carbon was already fixed by oxygenic photosynthesis, the CO2-fixation of anoxygenic phototrophic bacteria does not lead to a net increase in organic carbon available to higher trophic levels. The CO2-assimilation by anoxygenic phototrophic bacteria has therefore been termed “secondary primary production” (Pfennig, 1978). Therefore, capture of light energy by anoxygenic photosynthesis merely compensates for the degradation of organic carbon in the anaerobic food chain. Geothermal sulfur springs are the only exception since their sulfide is of abiotic origin. However, because sulfur springs are rather scarce, anoxygenic photosynthetic carbon fixation of these ecosystems also appears to be of minor significance on a global scale. The scientific interest in anoxygenic phototropic bacteria stems from 1) the simple molecular architecture and variety of their photosystems, which makes anoxygenic phototrophic bacteria suitable models for biochemical and biophysical study of photosynthetic mechanisms, 2) the considerable diversity of anoxygenic phototrophic bacteria, which has implications for reconstructing the evolution of photosynthesis, and 3) the changes in biogeochemical cycles of carbon and sulfur, which are mediated by the dense populations of phototrophic bacteria in natural ecosystems. All known microorganisms use two functional principles (both mutually exclusive and represent two independent evolutionary developments) for the conversion of light into chemical energy. Chlorophyll-based systems are widespread among members of the domain Bacteria and consist of a light-harvesting antenna and reaction centers. In the latter, excitation energy is converted into a redox gradient across the membrane. In contrast, the retinal-based bacteriorhodopsin system is exclusively found in members of a monophyletic group within the domain Archaea. These prokaryotes lack an antenna system and use light energy for the direct translocation of protons across the cytoplasmic membrane. In both systems, photosynthetic energy conversion ultimately results in the formation of energy-rich chemical bonds of organic compounds. The advent of modern genetic and biochemical methods has led to a considerable gain in knowledge of the molecular biology of pho-

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totrophic prokaryotes. At the same time, microbial ecologists have found these microorganisms of considerable interest and now frequently use molecular methods to investigate natural populations. The present chapter is limited to the discussion of phototrophic bacteria and attempts to link the physiology, ecology, and evolution of phototrophic bacteria to a molecular basis. Emphasis is laid on those molecular structures or functions that have evident adaptive value. This integrating view may provide a more solid foundation for understanding the biology of photosynthetic prokaryotes.

Taxonomy of Phototrophic Prokaryotes The capacity for chlorophyll-based photosynthetic energy conversion is found in five of the 36 currently recognized bacterial lineages (Fig. 1; Hugenholtz et al., 1998): the Chloroflexus subgroup, the green sulfur bacteria, the Proteobacteria, the Cyanobacteria, and the Heliobacteriaceae. With the exception of the Cyanobacteria, phototrophic bacteria perform anoxygenic photosynthesis, which is not accompanied by photochemical cleavage of water and therefore does not lead to the formation of molecular oxygen. Based on their phenotypic characters, anoxygenic phototrophic bacteria had been divided previously into the five families Rhodospirillaceae, Chromatiaceae, Ectothiorhodospiraceae, Chlorobiaceae, and Chloroflexaceae (Trüper and Pfennig, 1981). However, 16S rRNA oligonucleotide cataloguing and 16S rRNA sequence comparisons have reveale d that the Proteobacteria and the Chloroflexus-subgroup both contain nonphototrophic representatives (Woese, 1987; Fig. 1). Therefore the use of light as an energy source for growth is not limited to phylogenetically coherent groups of bacteria. However, nonphototrophic representatives of the green sulfur bacterial and the cyanobacterial lineages have not been isolated to date. Within the Chloroflexus-subgroup, three different species (Chloroflexus aurantiacus, Chloroflexus aggregans and Heliothrix oregonensis) of filamentous multicellular phototrophs have been described. All three are thermophilic and grow photoorganoheterotrophically. In addition four mesophilic species (Oscillochloris chrysea, Oscillochloris trichoides, Chloronema giganteum, Chloronema spiroideum) have been affiliated with the Chloroflexus-subgroup based on their multicellular filaments, gliding motility, and the presence of chlorosomes containing bacteriochlorophylls c or d (Pfennig and Trüper, 1989). The phylogenetic position of these latter bacteria

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Fig. 1. Phylogenetic tree based on 16S rRNA sequences. All bacterial divisions containing culturable representatives were included in the analyses so that the phototrophic nature of the bacterial strains could be confirmed. Alignments were obtained with CLUSTAL W and pairwise distances calculated with the algorithm of Jukes and Cantor using the DNADIST program of PHYLIP 3.57c. The tree was constructed from evolutionary distances employing the least-squares algorithm of Fitch and Margoliash as implemented by the FITCH program of the package. The Archaeon Methanopyrus kandleri DSM 6324 was used as an outgroup to root the tree. (light green) Bacteria containing chlorosomes as light-harvesting antenna. (red) Bacteria containing antenna complexes within the cytoplasmic membrane and quinone/pheophytin-type reaction centers. (medium green) Gram-positive phototrophic bacteria with FeS-type reaction centers. (dark green) Bacteria containing the two types of reaction centers. Width of colored wedges indicates the phylogenetic divergence.

has not been investigated so far. With the exception of Heliothrix oregonensis all species mentioned contain chlorosomes as distinct lightharvesting structures (Fig. 2). Yet to be cultivated axenically, non-thermophilic “Chloroflexus-like” organisms are known from intertidal and hypersaline benthic environments (Pierson et al., 1994) and from cold freshwater sulfidic springs (F. Garcia-Pichel, unpublished observation). At least in the case of the hypersaline enrichments, the organisms are closely related to Heliothrix in terms of their 16S rRNA sequence (B.K. Pierson, personal communication to FGP). This, together with recent descriptions of Oscillochloris trichoides (Keppen et al., 1994) from freshwater sediments indicates a larger diversity and more widespread occurrence of the Chloroflexaceae and allied organisms than was previously recognized. Green sulfur bacteria (see The Family Chlorobiaceae Volume 7) represent a coherent and isolated group within the domain Bacteria. They are strict photolithotrophs and contain chlorosomes (Fig. 3A). During the oxidation of sulfide, elemental sulfur is deposited extracellularly. Another typical feature of this group is the very limited physiological flexibility (see Docile Reac-

tion). In the Proteobacteria, the a- and b-Proteobacteria comprise photosynthetic representatives (often also called the purple nonsulfur bacteria), which do not form separate phylogenetic clusters but are highly intermixed with various other phenotypes. Characteristically, members of these two groups exhibit a high metabolic versatility and are capable of photoorganotrophic, photolithoautotrophic and chemoorganotrophic growth. Photosynthetic pigments are bacteriochlorophyll a or b and a variety of carotenoids. Light-harvesting complexes, reaction centers, and the component s of the electron transport chain are located in intracellular membrane systems of species-specific architecture (Fig. 2; see Light Absorption and Light Energy Transfer in Prokaryotes in this Chapter). Several members of the a-Proteobacteria are capable of bacteriochlorophyll a synthesis but cannot grow by anoxygenic photosynthesis. This physiological group has therefore been designated “aerobic anoxygenic phototrophic bacteria” (Shimada, 1995; Yurkov and Beatty, 1998), “aerobic phototrophic bacteria” (Shiba, 1989), or “quasi-photosynthetic bacteria” (Gest, 1993) and comprises a considerable number of species. So far, the marine genera Erythrobacter and

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Fig. 2. Organization of the phototrophic apparatus in different groups of phototrophic bacteria. OM = outer membrane, CW = cell wall, CM = cytoplasmic membrane, RC = reaction center, LHC = lightharvesting complex. Question marks indicate that the organization of the cell envelope and the organization of the photosynthetic apparatus in Heliothrix oregonensis is not exactly known.

Roseobacter and the six freshwater genera Acidiphilium, Erythromonas, Erythromicrobium, Porphyrobacter, Roseococcus, Sandarcinobacter (Yurkov and Beatty, 1998) have been described. This group also includes some aerobic facultatively methylotrophic bacteria of the genus Methylobacterium and a Rhizobium (strain BTAi1; Evans et al., 1990; Shimada, 1995; Urakami and Komagata, 1984). The oxidation of organic carbon compounds is the principal source of metabolic energy. Photophosphorylation can be used as a supplementary source of energy, with a transient enhancement of aerobic growth following a shift from dark to illumination (Harashima et al., 1978; Shiba and Harashima, 1986). Aerobic bacteriochlorophyllcontaining bacteria harbor a photosynthetic apparatus very similar to photosystem II of anoxygenic phototrophic Proteobacteria

(Yurkov and Beatty, 1998). Photochemically acti ve reaction centers and light-harvesting complexes are present, as are the components of cyclic electron transport (e.g., a cytochrome c bound to the reaction center and soluble cytochrome c2). In contrast to anoxygenic phototrophic bacteria, however, the aerobic phototrophic bacteria cannot grow autotrophically. Intracellular photosynthetic membrane systems as they are typical for anoxygenic phototrophic Proteobacteria are absent in most aerobic photosynthetic bacteria; Rhizobium BTAi1 being a possible exception (Fleischman et al., 1995). The presence of highly polar carotenoid sulfates and C30 carotenoid glycosides is a unique property of this group. All aerobic bacteriochlorophyll a-containing species group with the a-subclass of the Proteobacteria, but are more closely related to aerobic non-

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bacteriochlorophyll-contain ing organisms than to anoxygenic phototrophs (Stackebrandt et al., 1996). The g-subclass comprises two families of phototrophic species, the Chromatiaceae and Ectothiorhodospiraceae (also called purple sul-

CHAPTER 1.3

fur bacteria). Chromatiaceae accumulate sulfur globules within the cells and represent a conspicuous microscopic feature of these bacteria. With one notable exception (Thiocapsa pfennigii), the intracellular membrane system is of the vesicular type (Figs. 2 and 3B). In contrast, members of

Fig. 3. Localization and organization of the photosynthetic apparatus in three major groups of phototrophic bacteria. Electron-donating enzyme systems, like flavocytochrome or sulfide quinone reductase, and ATP formation by the membranebound ATP synthase are not shown. A. Green sulfur bacteria (Chlorobiaceae). B. Purple nonsulfur bacteria and Chromatiaceae. C. Cyanobacteria. OM = outer membrane; CW = cell wall; CM = cytoplasmic membrane; Cyt = cytochrome; P840 and P870 reaction center special pair = primary electron donor; B800, B850, B875 = bacteriochlorophyll molecules bound to lightharvesting complexes II and I; A0 = primary electron acceptor in green sulfur bacteria = Chl a; A1 = secondary electron acceptor in green sulfur bacteria = menaquinone; QA, QB = ubiquinone; FX, FA, FB = FeS-clusters bound to the reaction center; Fd = ferredoxin; FMO = Fenna-Matthews-Olson protein; FNR = ferredoxin NADP+ reductase; PQ = plastoquinone; PC = plastocyanin; PS = photosystem.

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Fig. 3. Continued.

the Ectothiorhodospiraceae deposit elemental sulfur outside of the cells and contain lamellar intracellular membrane systems. Like their relatives of the a- and b-subclass of Proteobacteria, the purple sulfur bacteria contain bacteriochlorophylls a and b, and all components of the photosynthetic apparatus are located in the intracellular membrane. No photosynthetic species have been described for the d- or e-subclass of the Proteobacteria. Heliobacteriaceae differ from other anoxygenic phototrophic bacteria by their unique light-harvesting and reaction center pigment,

bacteriochlorophyll g, and by their phylogenetic affiliation (Fig. 1). The first member of this group, Heliobacterium chlorum was described in 1983 by Gest and Favinger (Gest and Favinger, 1983b). Based on peptidoglycan structure studies (Beer-Romero et al., 1988), their high proportion of branched-chain fatty acids (Beck et al., 1990) and 16S rRNA sequencing, the Heliobacteriaceae belong to the Gram-positive low GC lineage. A close relatedness can also be deduced from the capability of Heliobacterium modesticaldum and Heliobacterium gestii to form endospores. However, a detailed phylogenetic analysis also indicated a close relatedness of

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CHAPTER 1.3

Fig. 3. Continued.

Heliobact eriaceae to the Cyanobacteria (Vermaas, 1994). Heliobacteriaceae do not contain distinct intracellular structures of the photosynthetic apparatus and the reaction centers are located in the cytoplasmic membrane. Bacteriochlorophyll g confers to the cells a near infrared absorption maximum at 788 nm, which is unique among photosynthetic organisms. The known species of Heliobacteriaceae all grow photoheterotrophically and are strict anaerobes. Oxygenic photosynthesis is only found in members of a single bacterial lineage out of the

five that contain phototrophs (Fig. 1). The Cyanobacteria by far comprise the largest number of isolated strains and described species (Table 1). The Cyanobacteria (= oxyphotobacteria) are defined by their ability to carry out oxygenic photosynthesis (water-oxidizing, oxygen-evolving, plant-like photosynthesis) based on the coordinated work of two photosystems (Fig. 3C). Phylogenetically, they constitute a coherent phylum that contains the plastids of all eukaryotic phototrophs. They all synthesize chlorophyll a as photosynthetic pigment, and

CHAPTER 1.3

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Table 1. Groups of photosynthetic prokaryotes and their characteristics. Taxon Chloroflexus subdivision

Preferred growth mode (3)a

Light harvesting

Anoxygenic photoorganoheterotroph(cls); Aerobic chemoorganoheterotroph

BChl c, car

Photochemical reaction Type II reaction center

—

—

Green sulfur bacteria

(15)

Anoxygenic photolithoautotroph

cls; BChl cldle, car

Type I reaction center

a-Proteobacteria

(31)

Anoxygenic photoorganoheterotroph Aerobic chemoorganoheterotroph

icm; BChl alb, car

Type II reaction center

—

—

a-Proteobacteria (aerobic photosynthetic)

(23)

Aerobic chemoorganoheterotroph

BChl a

Type II reaction center

b-Proteobacteria

(4)

Anoxygenic photoorganoheterotroph Aerobic chemoorganoheterotroph

icm; BChl a, car

Type II reaction center

—

—

Chromatiaceae Ectothiorhodospiraceae

(31) (9)

Anoxygenic photolithoautotroph

icm; BChl alb, car

Type II reaction center

Heliobacteriaceae

(5)

Anoxygenic photoorganoheterotroph

BChl g, car

Type I reaction center

Cyanobacteria

(>> 1000)

Oxygenic photolithoautotroph

thy; Chl a + PBS or Chl b, or Chl d; car

Type I + II reaction center

Prochloron, Prochlorothrix

(2)

thy; Chl a/b,car

Prochlorococcus

(1)

thy; Chl a2/b2, car (PBS)

Acaryochloris

(1)

thy; Chla,d, car (PBS)

Halobacteria

(3)

Aerobic chemoorganoheterotroph

Purple membrane; bacteriorhodopsin

Bacteriorhodopsin

a

The numbers of photosynthetic species described for each taxon are given in parenthesis. BChl = bacteriochlorophyll, car = carotenoids, Chl = chlorophyll, cls = chlorosomes, icm = intracellular membranes, PBS = phycobilisomes, thy = thylacoids.

most types contain phycobiliproteins as lightharvesting pigments. These multimeric proteinaceous structures are found on the cytoplasmic face of the intracellular thylakoid membranes and contain phycobilins as light-harvesting pigments. All Cyanobacteria are able to grow using CO2 as the sole sou rce of carbon, which they fix using primarily the reductive pentose phosphate pathway (see Carbon Metabolism of Phototrophic Prokaryotes in this Chapter). Their chemoorganotrophic potential typically is restricted to the mobilization of reserve polymers (mainly starch but also polyhydroxyalkanoates) during dark periods, although some strains are known to grow chemoorganotrophically in the dark at the expense of external sugars. Owing to their ecological role, in many cases indistinguishable from that of eukaryotic microalgae, the cyanobacteria had been studied originally by botanists. The epithets “blue-green algae,” “Cyanophyceae,” “Cyanophyta,” “Myxophyceae,” and “Schizophyceae” all apply to the cyanobacteria. Two main taxonomic treatments of the Cyanobacteria exist, and are widely used, which divide them into major groups (orders) on

the basis of morphological and life-history traits. The botanical system (Geitler, 1932 recognized 3 orders, 145 genera and some 1300 spe cies, but it has recently been modernized (Anagnostidis and Komárek, 1989, Komárek and Anagnostidis, 1989). The bacteriological system (Stanier, 1977; Rippka et al., 1979; Castenholz, 1989), relies on the study of cultured axenic strains. It recognizes five larger groups or orders, separated on the basis of morphological characters. Genetic (i.e., mol% GC, DNA-DNA hybridization) as well as physiological traits have been used to separate genera in problematic cases. Previously, a separate group of organisms with equal rank to the cyanobacteria, the so-called “Prochlorophytes” (with two genera, Prochloron, a unicellular symbiont of marine invertebrates, and Prochlorothrix, a free-living filamentous form) had been recognized (Lewin, 1981). They were differentiated from cyanobacteria by their lack of phycobiliproteins (Fig. 2) and the presence of chlorophyll b. The recently recognized genus Prochlorococcus of marine picoplankters could be included here, even though the major chlorophylls in this genus are

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divinyl-Chl a and divinyl-Chl b. Fourteen Prochloron isolates from different localities and hosts have been found to belong to a single species by DNA-DNA hybridization studies (Stam et al., 1985; Holtin et al., 1990). Some of the original distinctions leading to the separation of the Chl b-containing oxyphotobacteria from the cyanobacteria are questionable, since at least in one strain of Prochloroccoccus marinus, functional phycoerythrin (Lokstein et al., 1999), and genes encoding for phycobiliproteins have been detected (Lokstein et al., 1999). Additionally, phylogenetic analysis of 16S rRNA genes indicate that the three genera of Chl b-containing prokaryotes arose independently from each other and from the main plastid line (see Evolutionary Considerations in this Chapter), a result that is supported by the comparative sequence analysis of the respective Chl a/b binding proteins (Laroche et al., 1996; Vanders taay et al., 1998). Thus “Prochlorophytes” are just greenish cyanobacteria, and are not treated separately here. The recent discovery of Chl d-containing symbionts in ascidians (Acaryochloris marina, Miyashita et al., 1996) once again demonstrates the evolutionary diversification of light-harvesting capabilities among oxyphotobacteria (see Competition for Light in this Chapter). While the phylogenetic affiliation of Acaryochloris marina has not been presented as yet, ultrastructural and chemotaxonomic characters predict that A. marina belongs to the cyanobacterial radiation as well. According to phylogenetic analysis of 16S rRNA sequences, the Cyanobacteria are a diverse phylum of organisms within the bacterial radiation, well separated from their closest relatives (Giovanonni, 1988; Wilmotte, 1995; Turner, 1887; Garcia-Pichel, 1999; Fig. 1). These analyses support clearly the endosymbiotic theory for the origin of plant chloroplasts, as they place plastids (from all eukaryotic algae and higher plants investigated) in a diverse, but monophyletic, deep-branching cluster (Nelissen et al., 1995). Phylogenetic reconstructions show that the present taxonomic treatments of the cyanobacteria diverge considerably from a natural system that reflects their evolutionary relationships. For example, separation of the orders Chroococcales and Oscillatoriales (Nelissen et al., 1995; Reeves, 1996), and perhaps also the Pleurocapsales (Turner, 1887; Garcia-Pichel et al., 1998) is not supported by phylogenetic analysis. The heterocystous cyanobacteria (comprising the two orders Nostocales and Stigonematales) form together a monophyletic group, with relatively low sequence divergence, as low as that presented by the single accepted genus Spirulina (Nübel, 1999). A grouping not corresponding to any official genus, the Halothece cluster, gathers

CHAPTER 1.3

unicellular strains of diverse morphology that are extremely tolerant to high salt and stem from hypersaline environments (Garcia-Pichel et al., 1998). A second grouping, bringing together very small unicellular ope n-ocean cyanobacteria (picoplankton) includes only marine picoplanktonic members of the genera Synechococcus and all Prochlorococcus. Several other statistically well-supported groups of strains that may or may not correspond to presently defined taxa can be distinguished. The botanical genus “Microcystis” of unicellular colonial freshwater plankton species is very well supported by phylogenetic reconstruction, as is the genus Trichodesmium of filamentous, nonheterocystous nitrogen-fixing species typical from oligotrophic marine plankton of the tropics. The picture that emerges from these studies is that sufficient knowledge of ecological and physiological characteristics can lead to a taxonomic system that is largely congruent to the 16S rRNA phylogeny. A different principle of conversion of light energy into chemical energy is found in the Halobacteria. These archaea are largely confined to surface layers of hypersaline aquatic environments and grow predominantly by chemoorganoheterotrophy with amino or organic acids as electron donors and carbon substrates, generating ATP by respiration of molecular oxygen. In the absence of oxygen, several members are capable of fermentation or nitrate respiration. At limiting concentrations of oxygen, at least three of the described species of Halobacteria (Halobacterium halobium, H. salinarium, H. sodomense) synthesize bacteriorhodopsin (Oesterhelt and Stoeckenius, 1973), a chromoprotein containing a covalently bound retinal. Bacteriorhodopsin is incorporated in discrete patches in the cytoplasmic membrane (“purple membrane”). However, these prokaryotes have only a very limite d capability of light-dependent growth. Only slow growth and one to two cell doublings could be demonstrated experimentally (Hartmann et al., 1980; Oesterhelt and Krippahl, 1983). The fact that rhodopsin-based photosynthesis has been found only in the phylogenetically tight group of Halobacteria may indicate that, because of its lower efficiency, this type of light utilization is of selective advantage only under specific (and extreme) environmental conditions. Further information on the biochemistry, physiology and ecology of this group may be found in the chapters, Introduction to the Classification of Archaea and The Family Halobacteriaceae. During the past years, culture-independent 16S rDNA-based methods have been used for the investigation of the composition of natural communities of phototrophic prokaryotes. These studies have provided evidence that more than

CHAPTER 1.3

one genotype of Chloroflexus occur in one hot spring microbial mat and that four previously unkown sequences of cyanobacteria dominate in the same environment (Ferris et al., 1996; RuffRoberts et al., 1994; Weller et al., 1992). Similarly, nine different partial 16S rDNA sequences of Chromatiaceae and green sulfur bacteria, which differed from all sequences previously known, were retrieved from two lakes and one intertidal marine sediment (Coolen and Overmann, 1998; Overmann et al., 1999a). However, 16S RNA signatures from natural populations were indistinguishable from those of cultured strains in the case of cyanobacteria with conspicuous morphologies, such as the cosmopolitan Microcoleus chthonoplastes (Garcia-Pichel et al., 1996) from intertidal and hypersaline microbial mats or Microcoleus vaginatus from desert soils (F. Garcia-Pichel, C. López-Cortés and U. Nübel, unpublished observations). In a similar manner, the 16S rRNA sequence of an isolated strain of Amoebobacter purpureus (Chromatiaceae) was found to be identical to the environmental sequence dominating in the chemocline of a meromictic salt lake (Coolen and Overmann, 1998; Overmann et al., 1999a). Obviously, the limited number of isolated and characterized bacterial strains rather than an alleged “nonculturability,” at least in some cases, accounts for our inability to assign ecophysiological properties to certain 16S rRNA sequence types. This point is illustrated for extremely halotolerant unicellular cyanobacteria by the fact that only after a physiologically coherent group of strains was defined on the basis of newly characterized isolates (Garcia-Pichel et al., 1998) could the molecular signatures retrieved from field samples be assigned correctly. It has to be concluded that 1) the numbers of species listed in Table 1 do not reflect the full phylogenetic breadth at least in the four groups of anoxygenic phototrophic prokaryotes as well as in morphologically simple Cyanobacteria, and 2) that the physiology and ecology of those species of phototrophic prokaryotes that are dominant in the natural environment in some cases may differ considerably from known type strains.

Habitats of Phototrophic Prokaryotes Bacteria of the Chloroflexus-subgroup form dense microbial mats in geothermal springs, often in close association with cyanobacteria. Chloroflexus aurantiacus is a thermophilic bacterium which grows optimally between 52 and 60∞C and thrives in neutral to alkaline hot springs up to 70–72∞C. Of all anoxygenic phototrophic bacteria isolated so far, only Chlorof-

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lexus aurantiacus is capable of growth up to 74∞C. In contrast to the domain Archaea, no hyperthermophilic species are known from the domain Bacteria. The phylogenetically related Heliothrix oregonensis grows optimally between 50 and 55∞C and is abundant as a flocculant surface layer in a few alkaline springs in Oregon. Hydrothermal springs of 56–66∞C, which contain sulfide of geothermal origin, are dominated by a surface layer or a “unispecific” mat of Chloroflexus (Castenholz and Pierson, 1995). Because of the absence of cyanobacteria in some of these systems, Chloroflexus presumably grows autotrophically (Pierson and Castenholz, 1995). In the presence of O2, the mats exhibit an orange color whereas they are green under anoxic conditions (Castenholz and Pierson, 1995). The orange color is the result of the enhanced carotenoid biosynthesis under oxic conditions (see Chemotrophic Growth with O2 in this Chapter). In the absence of sulfide, Chloroflexus is present as a distinct orange layer beneath a surface layer of cyanobacteria and may utilize their exudates or the fermentation products generated during decomposition of cyanobacteria. Molecular oxygen represses bacteriochlorophyll synthesis in Chloroflexus and often is present at saturation levels in the orange layers. Since bacteriochlorop hylls a and c are still present in this layer, however, it must be assumed that bacteriochlorophylls are synthesized at anoxic conditions during nightime (Castenholz and Pierson, 1995). Green and purple sulfur bacteria often form conspicuous blooms in non-thermal aquatic ecosystems (Figs. 4, 5A, 5B), although moderately

Fig. 4. Bright field photomicrograph of the bacterioplankton community thriving in the chemocline of the meromictic Buchensee (near Radolfzell, Germany) during autumn. The dominant anoxygenic phototroph at this time of the year is the green sulfur bacterium Pelodictyon phaeoclathratiforme (brown cells, which appear in chains or netlike colonies). In addition, phototrophic consortia (“Pelochromatium roseum,” one consortium in the center) are found. Similar to Pld. phaeoclathratiforme, most of the colorless bacterial cells found in the chemocline contain gas vesicles as is evident from their highly refractile appearance in the bright field.

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Fig. 5. Multilayered microbial mat as it is regularly found in the sandflats of Great Sippewissett Salt Marsh (Cape Cod, Massachusetts, USA). A. In most instances, the mats consist of a top green layer, an intermediate purple layer, and a grayish to blackish bottom layer. B. Fully developed microbial mats consist (from top) of an olive-green layer of diatoms and cyanobacteria, a green layer consisting mostly of cyanobacteria, a purple layer of purple sulfur bacteria, a peach-colored layer formed by BChl b-containing purple sulfur bacteria (morphologically similar to Thiocapsa pfennigii), and a greyish to blackish bottom layer.

thermophilic members of the genera Chromatium and Chlorobium have been described from hot spring mats (Castenholz et al., 1990). Chlorobium tepidum occurs in only a few New Zealand hot springs at pH values of 4.3 and 6.2 and temperatures up to 56∞C. Chromatium tepidum was found in several hot springs of western North America at temperatures up to 58∞C and might represent the most thermophilic proteobacterium (Castenholz and Pierson, 1995). In a recent compilation (van Gemerden and Mas, 1995), 63 different lakes and 7 sediment ecosystems harboring phototrophic sulfur bacteria were listed. Cell densities between 104 and 107·ml-1 and biomass concentrations between 10 and 1000 mg bacteriochlorophyll·l-1 are common in pelagic habitats. Of the purple sulfur bacteria, Chromatiaceae are typically found in freshwater and marine environments (Fig. 5A, B) whereas Ectothiorhodospiraceae inhabit hypersaline waters. The phototrophic sulfur bacteria grow preferentially by photolithoautotrophic oxida-

tion of reduced sulfur compounds and are therefore limited to those environments where light reaches anoxic, sulfide-containing bottom layers. Because light and sulfide occur in opposing gradients, growth of phototrophic sulfur bacteria is confined to a narrow zone of overlap and is only possible if the chemical gradient of sulfide is stabilized against vertical mixing. In pelagic environments like lakes or lagoons, chemical gradients are stabilized by density differences between the oxic and anoxic water layers. Such density differences are either the result of thermals tratification and mostly transient (as in holomictic lakes) or are caused by high salt concentrations of the bottom water layers, in which case stratification is permanent (meromictic lakes). Pelagic layers of phototrophic sulfur bacteria extend over a vertical distance of 10 cm (van Gemerden and Mas, 1995; Overmann et al., 1991a) up to 30 m (Repeta et al., 1989) and reach biomass concentrations of 28 mg bacteriochlorophyll·l-1 (Overmann et al., 1994).

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43

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The Phototrophic Way of Life 400

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CHAPTER 1.3

Wavelength (nm) Fig. 6. Effects of the habitat on the physical exposure of cyanobacteria. The spectral scalar irradiance (sun and sky radiation) incident at ground level at noon in a clear midsummer day at 41∞N is plotted in Plate I. The rest of the plates depict the in situ scalar irradiance experienced by cyanobacterial cells thriving in several habitats exposed to the incident fluxes in plate I (note different scales). Plate II: a “strong shade” habitat (North-facing surface illuminated by extremely diffuse sky radiation only), where scalar irradiance is very low but the relative importance of UV is enhanced. Plate III: a planktonic habitat (under 1 m of clear open-ocean water), where all fluxes remain fairly high and UVB and visible are more strongly attenuated than UVA. Plate IV: the surface of beach (quartz, feldspar) sand, where all UVB, UVA, and visible are higher than incident (by 120, 150, and 205%, respectively) due to light trapping effects. Plate V: 300-m deep in a wet topsoil, where UVB and U VA have been attenuated below 5% of incident but ca. 20% of the visible light remains. Plate VI: scalar irradiance within the thallus of the terrestrial cyanobacterial lichen Collema sp. Modified from Castenholz and Garcia-Pichel, 1999, after data from the following sources: F. Garcia-Pichel (unpublished observation); Garcia-Pichel, 1995; Büdel et al., 1997; and Smith and Baker, 1981.

Littoral sediments represent the second type of habitat of phototrophic sulfur bacteria. In these systems, turbulent mixing is largely prevented by the sediment matrix, and diffusion is the only means of mass transport. Gradients of light and sulfide are much steeper, and the fluxes of sulfide much larger compared to the pelagic environment. These conditions allow layers of phototrophic sulfur bacteria in sediments to reach much higher biomass densities (up to 900 mg bacteriochlorophyll·dm-3; van Gemerden et al., 1989) than in lakes. At the same time, the layers are very narrow (1.3– 5 mm; van Gemerden and Mas, 1995; Fig. 5A). This vertical distribution of anoxygenic phototrophic biomass ultimately determines the significance of microbial sulfide oxidation for the sulfur cycle in these ecosystems (see Significance of Anoxygenic Photosynthesis for the Pelagic Carbon and Sulfur Cycles in this Chapter). The spectral compos ition of light available for anoxygenic photosynthesis is considerably different between pelagic and benthic habitats (Fig. 6) and selects for different species of anoxygenic phototrophic bacteria. Whereas light of the blue to yellow-green wavelength bands dominates the depths of most lakes, infrared light is an important source of energy in benthic microbial mats (see Light Energy

and the Spectral Distribution of Radiation in this Chapter). The dominance of certain species of green sulfur bacteria (Fig. 4) or Chromatiaceae in pelagic environments in many cases can be explained by their specific light-harvesting capabilities (see Light Absorption and Light Energy Transfer in Prokaryotes and Competition for Light in this Chapter) and other phenotypic traits. Typically, those species that have been isolated from natural blooms in lakes are obligately photolithotrophic, lack assimilatory sulfate reduction, cannot reduce nitrate, and assimilate only few organic carbon sources (see Carbon Metabolism of Phototrophic Prokaryotes in this Chapter). This applies not only to all green sulfur bacteria but also to the dominant species of Chromatiaceae. Obviously, in the chemocline of lakes the metabolic versatile Chromatiaceae species have no selective advantage. As judged from the physiological characteristics of strains of phototrophic sulfur bacteria isolated from sediments, the pronounced diurnal variations in oxygen concentrations and salinity, together with the different light quality, select for different species composition in benthic microbial mats. The purple sulfur bacterium Chromatium (and the multicellular gliding colorless sulfur bacterium Beggiatoa) are found in many microbial mats

44

J. Overmann and F. Garcia-Pichel

and exhibit diurnal vertical migrations in response to the recurrent changes in environmental conditions (Jørgensen, 1982; Jørgensen and Des Marais, 1986). Microbial mats of intertidal sediments are typically colonized by the immotile purple sulfur bacterium Thiocapsa roseopersicina and small motile thiobacilli (van den Ende et al., 1996). In contrast to the phototrophic members of the g-Proteobacteria, purple nonsulfur bacteria of the a- and b-subclasses of Proteobacteria do not appear to form dense accumulations under natural conditions (Biebl and Drews, 1969; Swoager and Lindstrom, 1971; Steenbergen and Korthals, 1982). However, purple nonsulfur bacteria can be readily isolated from a wide variety of marine, lacustrine and even terrestrial environments (Imhoff and Trüper, 1989; J. Overmann, unpublished observation). While comprehensive comparative quantitation of the ecological importance of purple nonsulfur bacteria is still lacking, as many as ca. 106 c.f.u. of purple nonsulfur bacteria could be cultivated per cm3 of sediment in coastal eutrophic settings (Guyoneaud et al., 1996). Generally, aerobic phototrophic bacteria thrive in eutrophic marine environments. Obligately aerobic bacteria containing bacteriochlorophyll a have been isolated from beach sand and seaweeds (thalli of Enteromorpha linza and Sargassum horneri; Shiba et al., 1979), and in some cases also from freshwater ponds and microbial mats. At least some of the aerobic phototrophic bacteria apparently can survive in situ temperatures of up to 54∞C (Yurkov and Beatty, 1998). Aerobic phototrophic bacteria were isolated from hydrothermal plume water of a black smoker 2000 m below ocean surface (Yurkov and Beatty, 1998); acidophilic strains could be isolated from acidic mine drainage. Typically, Methylobacterium species are isolated from foods, soils and leaf surfaces (Shimada, 1995). Photosynthetic Rhizobium strains are widely distributed in nitrogen-fixing stem nodules of the tropical legume Aeschynomene spp. where they are present as symbiosomes. Similar strains have also been found in root and hypocotyl nodules of Lotononis bainesii (Fabaceae). These photosynthetic rhizobial and regular symbiosomes differ in that the former contains only one large spherical bacteroid. The photosynthesis of these endosymbionts may provide energy for nitrogen fixation and permit a more efficient growth of the host plant, since up to half of the photosynthate produced by legumes is allocated to nitrogen fixation (Fleischman et al., 1995). Heliobacteriaceae appear to be primarily soil bacteria and have been isolated from dry paddy fields or other soils throughout the world (Madigan and Ormerod, 1995). Bacteria of this family

CHAPTER 1.3

may even represent the dominant anoxygenic phototrophic bacteria in soil (Madigan, 1992). Occasionally, strains also have been isolated from lakeshore muds and hot springs (Amesz, 1995; Madigan and Ormerod, 1995). Heliobacterium modesticaldum grows up to 56∞C (Kimble et al., 1995). Spore formation may offer a selective advantage to Heliobacterium modesticaldum, Heliophilum fasciatum, and Heliobacterium gestii in their main habitat (rice field soil), which undergoes periodic drying and concomitantly becomes oxidized (Madigan, 1992). During growth of the rice plants, organic compounds excreted by their roots could provide sufficient substrates for photoheterotrophic growth of the Heliobacteriaceae. Cyanobacteria as a group exhibit the widest range of habitats of all phototrophic prokaryotes due to the ubiquity of water, their preferred electron donor for the reduction of CO2. In principle, cyanobacteria can thrive in any environment that has, at least temporarily, liquid water and sunlight. They are known from Antarctic endolithic habitats and from hot springs. More than 20 species of cyanobacteria (Castenholz and Pierson, 1995) are thermophilic. Effectively, however, no cyanobacteria are known from acidic environments (below pH 4.5) and competition with eukaryotic microalgae or higher plants may restrict their growth in other environments. Cyanobacteria are found in the plankton of coastal and open oceans and in freshwater and saline inland lakes. They thrive in the benthos of marine intertidal (Fig. 5B), lacustrine and fluvial waters and in a large variety of terrestrial habitats (soils, rocks, trees). Symbiotic associations are common. In the marine plankton, the phycoerythrincontaining Synechococcus often represents a major fraction of all primary producers. The same holds true for Prochlorococcus (Campbell and Vaulot, 1993; Chisholm et al., 1988; Olson et al., 1990b). Compared with the high number of cyanobacterial species found in freshwater plankton, intertidal areas, and hypersaline environments, the diversity of this group is very limited in the open ocean (Carr and Mann, 1994). The predominant group invariably consists of small (97%) serves in light-harvesting and transfers excitation energy to the photochemical reaction centers. The combination of antenna complexes with one reaction center constitutes the photosynthetic unit. The efficiency of energy transfer within the photosynthetic unit and its size determine the fraction of the quantum flux that is harvested. Large concentrations of pigments result in self-shading and thus a reduced efficiency of light absorption per mole of pigment. At the cell size and intracellular pigment concentrations typical of most prokaryotic phototrophs, this decrease in efficiency is not very important (Garcia-Pichel, 1994a), but it might be significant in some extremely low-light adapted anoxygenic phototrophs like the green sulfur bacterial strain isolated from the Black Sea chemocline (Overmann et al., 1991a). Close proximity of photosynthetic pigments enables an efficient transfer of excitation energy but at the same time also causes a so-called “package effect” (Kirk, 1983) by which selfshading of the pigment molecules exceeds that predicted by the Lambert-Beer law. The package effect is seen clearly in a flattening of absorption peaks, commonly observed when recording absorption spectra of whole cells (see The Family Chlorobiaceae, Identification section in Volume 7). Because the energy requirement for biosynthesis of additional antenna structures is rather constant, the net energy gain for a photosynthetic cell must decrease at higher intracellular pigment concentrations, which restricts the

CHAPTER 1.3

The Phototrophic Way of Life

amount of light-harvesting structures a photosynthetic cell can synthesize. Polypeptides of the photosynthetic machinery (a significant fraction of the total cell protein) amount to 20% in purple nonsulfur bacteria and >50% in phycobiliprotein-containing cyanobacteria. Interestingly, the total protein content of cyanobacterial cells is comparable to other phototrophic bacteria. Possibly, cyanobacteria contain reduced levels of proteins involved in nonphotosynthetic processes to compensate for the high energy and nitrogen expenditure of the antenna proteins. The biosynthesis of proteins requires a major fraction of the energy expenditure of the bacterial cell (Gottschalk, 1986). In chlorosomes, the mass ratio of protein:bacteriochlorophyll is significantly lower than in other light-harvesting complexes (Table 3). Probably this is one major reason for the larger antenna size and the lower light energy requirements of green sulfur bacteria as compared to their purple and cyanobacterial counterparts (see Competition between Phototrophic Bacteria in this Chapter), and might help explain the competitive advantage gained by Prochlorocococcus over their close relatives Synechococcus in the open oceans.

Conversion of Light into Chemical Energy PRINCIPLE The unifying principle of bacterial and archaeal photosynthesis is the light-driven generation of a proton-motive force (PMF). The PMF is subsequently used by ATP synthase to form ATP, or for active transport and motility. In chlorophyll-based photosynthesis, redox reactions and charge separation precede the establishment of the PMF. In addition, reducing Table 3. Pigment:protein ratio in different photosynthetic antenna complexes.

51

power (NAD(P)H + H+) is generated as a primary product of the light reaction in Cyanobacteria. In the photochemical reaction, only the energy of the lowest excited singlet state (see Light Absorption and Light Energy Transfer in Prokaryotes in this Chapter) of the chlorophylls is used. Consequently, all absorbed light quanta have the same effect irrespective of their original energy (wavelength). When comparing the light energy available in different habitats, or the light adaptation of different phototrophic bacteria, it is therefore more meaningful to express irradiances in units of mol quanta·m-2·s-1 rather than W·m-2 (see Competition for Light in this Chapter). The standard free energy for the reduction of CO2 depends on the redox potential of the photosynthetic electron donor employed (Table 4, Fig. 8). If this energy requirement for electron transfer is compared with the energy available after absorption of photons of different wavelengths, it becomes clear that oxygenic photosynthesis is not feasible in photosystems containing the known types of chlorin pigments, and requires the absorption of two photons per electron (Fig. 8). The biological conversion of light into chemical energy has been found to be remarkably efficient: the number of charge separation events per absorbed photon is 1.0 (Kok, 1973; Wraight and Clayton, 1973) and the efficiency of the entire photoconversion process of a red photon to chemical energy by oxygenic photosynthetic organisms is 43% (Golbeck, 1994). Whereas the efficiency of energy transfer between antenna bacteriochlorophyll and the reaction center in most cases is close to 100% (Amesz, 1995), the transfer between antenna carotenoids and the reaction center can be significantly lower, 70% in Heliobacteriaceae (Amesz, 1995) and even

Protein:pigment Antenna complex type Chlorosomes B806-866 complexa B800-850 LHII B820 LHI Phycobilisomes

Mass ratio

Per pigment molecule (in Da)

0.5–2.2 3.9–5.8 4.4 6.7 ~22.4

420–1,840 3,550–5,290 4,000 6,100 ~12,300

a Chloroflexus aurantiacus. Data from Olson, 1998 or calculated from Sidler, 1994, Loach and Parkes-Loach, 1995, Zuber and Cogdell, 1995. Carotenoids have been neglected in these calculations because of their lower numbers as compared to bacteriochlorophylls (B800-850 LHII), their absence in phycobilisomes, and the controversy concerning their functional significance in lightharvesting (chlorosomes). Only antenna complexes which are separate entities from reaction centers were considered. Photosystem I does not contain a distinct antenna structure; the PsaA protein of the reaction center binds 110 chlorophyll a molecules.

Table 4. Standard redox potentials of different electron donors of the photosynthetic light reaction.a Electron donor 1

/2O2/H2O Fe(OH)3 + HCO3-/FeCO3 Fumarate/Succinate HSO3/S0SO42-/S0 SO42-/HSFe(OH)3/Fe2S0/HSHCO3-/acetate S2O32-/HS- + HSO3H+/ 1/2H2 Electron acceptor CO2/

Eo-[mV] +820 +200 +33 -38 -200 -218 -236 -278 -350 -402 -414 Eo-[mV] -434

a Taken from Brune, 1989; Widdel et al., 1993; Thauer et al., 1977; Zehnder and Stumm, 1988.

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CHAPTER 1.3

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Fig. 8. Free energy of one mol quanta calculated from Planck’s constant h (6.63 ¥ 1034 J·s), the speed of light c (2.99 ¥ 108 m·s1 ), the wavelengths of light l, and the Avogadro constant NA= 6.023 ¥ 1023 mol-1 according to DGóhn; = NA·h·c·l-1. Free energy required for the transfer of 1 mole of electrons from an electron donor with standard redox potential Eód (see Table 4) to CO2 calculated according to DGóel = F·(-470 - Eód) using the Faraday constant F (96.5 kJ·V-1·mol-1). Dotted vertical lines indicate the energy that is available after absorption of light by the long wavelength Qy absorption bands of different photosynthetic pigments.

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20% in a purple nonsulfur bacterium (Angerhofer et al., 1986). When carotenoids serve as the only light-harvesting pigments, 2.5 times higher irradiances are required by Rhodopseudomonas acidophila to attain the same growth rates as compared to light-absorption by bacteriochlorophyll (Göbel, 1978). In aerobic phototrophic bacteria, most of the highly diverse carotenoids do not function as light-harvesting molecules but might serve in quenching of toxic oxygen radicals (Noguchi et al., 1992; Yurkov et al., 1994). The same has been proposed recently for the carotenoid isorenieratene/b-isorenieratene in browncolored green sulfur bacteria (J. B. Arellano, J. Psencik, C. M. Borrego, R. Guyoneaud, C. A. Abella, L. J. Garcia-Gil, T. Gillbro, personal communication). One prerequisite for the photoconversion process is the presence of a membrane that is impermeable to protons and separates two different cell compartments. Three integral membrane multisubunit protein complexes participate in the generation of ATP in all phototrophic bacteria: the photosynthetic reaction center, a cytochrome complex, and an ATP synthase. All three are highly conserved within the bacterial radiation. Reaction centers have a dimeric core and consist of two closely associated integral mem-

E° (mV)

brane polypeptides plus additional proteins (Fig. 3). The special protein environment of the reaction center stabilizes the excited state and prevents back reaction after charge separation by enforcing ultrafast electron transfer to other electron acceptors nearby. The transfer of excitation energy from the antenna complexes to the reaction center initiates a charge separation at a special bacteriochlorophyll dimer (special pair), which is located on the periplasmic (or lumen) side of the photosynthetic mem brane. It is this endergonic process of charge separation that is ultimately driven by light energy; all the following redox reactions are exergonic. An electric potential is established across the membrane (inside negative). In its excited state, the special pair becomes a powerful reductant and ultimately reduces a quinone (in pheophytin-type reaction centers) or ferredoxin (in FeS-type reaction centers) on the cytoplasmic side of the photosynthetic membrane. The quinol or reduced ferredoxin leaves the reaction center complex and in turn donates electrons to a membranebound cytochrome complex or NADH dehydrogenase. A series of redox reactions results in the establishment of a proton-motive force across the photosynthetic membrane. Finally, the PMF is converted to ATP by ATPase.

CHAPTER 1.3

In contrast to the (bacterio)chlorophyll-based systems of bacteria, light energy conversion of Halobacteria does not involve redox reactions and is limited to a vectorial transport of protons by bacteriorhodopsin. Upon excitation by light, the prosthetic retinal undergoes a series of reversible photochemical transformations (an isomerization from the all-trans to the 13-cis form) and releases a proton into the extracellular space. The PMF thus generated is used for ATP synthesis by ATPase. Due to its low solubility, O2 in the concentrated salt solution is present in significantly lower amount than in freshwater. Rhodopsin-mediated formation of ATP may become the sole source of energy for growth under anaerobic conditions in the light (Oesterhelt and Krippahl, 1983) and has therefore been viewed as an adaptation to the natural brine habitat of Halobacteria. Because of its distinct mechanism, archaeal “photosynthesis” is not discussed in further detail in the present section. Additional information can be found in chapters titled Introduction to the Classification of Archaea and The Family Halobacteriaceae.

Molecular Architecture of the Reaction Center All bacteria which perform anoxygenic photosynthesis possess—or (in the case of cyanobacteria which are capable of using sulfide as electron donor) employ—only a single photosystem. The decrease in redox potential that a single photosystem can undergo upon excitation appears to be limited (Blankenship, 1992, compare Fig. 8). A combination of two different photosystems is required for the thermodynamically unfavorable utilization of water as an electron donor for photosynthesis (Fig. 3C). With the relatively simple architecture of their photosystems, all anoxygenic phototrophic bacteria depend on electron donors that exhibit standard redox potentials more negative than water (e.g., H2S, H2, acetate; Table 4). This molecular feature is one major reason for the narrow ecological niche of anoxygenic phototrophic bacteria in extant ecosystems (see Habitats of Phototrophic Prokaryotes in this Chapter). Two different types of reaction centers occur in photosynthetic bacteria. Based on the chemical nature of the early electron acceptors, a pheophytin/quinone-type reaction center and a FeS-type reaction center are distinguished (Blankenship, 1992; Fig. 3A,B). The first type is found in green gliding Chloroflexus species, phototrophic members of the a- and b-Proteobacteria, Chromatiaceae, Ectothiorhodospiraceae, and in PSII of Cyanobacteria. The reaction center of Proteobacteria consists of three protein subunits (L, M, H) which bind four

The Phototrophic Way of Life

53

bacteriochlorophylls, two bacteriopheophytins, two quinones and one high-spin nonheme Fe2+ (Lancaster and Michel, 1996; Fig. 3B). Many species (e.g., Chloroflexus aurantiacus, Blastochloris viridis and Allochromatium vinosum) contain an additional tetraheme cytochrome c polypeptide attached to the periplasmic side of the reaction center. Following the transfer of the electrons by ubiquinol or plastoquinol, the redox reactions at the cytochrome bc1 (or b6f) complex drive proton transport across the cytoplasmic membrane. Protons are translocated either into the extracellular space (anoxygenic phototrophic bacteria) or the intrathylacoidal space (cyanobacteria). The ratio of protons translocated to electrons transferred (H+/e- ratio) is 2. The reaction center and cytochrome bc1 in pheophytin-type reaction centers of Proteobacteria and Chloroflexus are functionally linked by two diffusible electron carriers, ubiquinone in the hydrophobic domain of the membrane and cytochrome c2 or auracyanin (Meyer and Donohue, 1995) in the periplasmic space. The liberated electron is transferred back to the special pair via quinone, the cytochrome bc1 complex and soluble periplasmic soluble electron carrie r (often cytochrome c2). Owing to this cyclic electron transport, the only primary product of photosynthesis is the proton-motive force, and the reduced pyridine nucleotide required for photosynthetic CO2 fixation is generated by energy-dependent reverse electron flow (Fig. 3). In oxygenic phototrophic bacteria, plastoquinone is the electron acceptor of PSII and donates electrons to the cytochrome b6fcomplex. The special pair is reduced by the manganese-containing water-splitting system located at the lumenal side of the transmembrane PSII complex (Fig. 3C). In the pheophytin-type reaction centers of aerobic phototrophic bacteria, photoinduced charge separation occurs only in the presence of O2 (Okamura et al., 1985). It has been proposed (Yurkov and Beatty, 1998) that oxic conditions are required for photochemical activity because the primary acceptor ubiquinone has a significantly higher midpoint redox potential than in anoxygenic photosynthetic bacteria (65 to 120 mV more positive). The primary acceptor therefore may stay in its oxidized, electronaccepting state only in the presence of O2. The second type of reaction center contains iron-sulfur clusters as early electron acceptors and occurs in green sulfur bacteria (Fig. 3A), Heliobacteriaceae, and in the photosystem I of Cyanobacteria. Functionally, the reaction centers of green sulfur bacteria, Heliobacteriaceae, and PSI of cyanobacteria are therefore similar. However, the former two are homodimeric and only

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J. Overmann and F. Garcia-Pichel

one reaction center gene has been detected, whereas the reaction center of PSI of cyanobacteria and green plants contains two nonidentical, but similar, subunits (PS I-A and PS I-B; Vermaas, 1994). In FeS-type reaction centers, the redox potential of the special pair in its reduced state (P*) is sufficiently low to permit a transfer of electrons to ferredoxin. Until recently, it has therefore been assumed that noncyclic electron flow can directly reduce NAD(P)+ and does not require further energy expenditure not only in cyanobacteria but also in green sulfur bacteria. However, the sequencing of the whole genome of Chlorobium tepidum has not provided any indications for the presence of a ferredoxinNADP+ oxidoreductase (D. A. Bryant, personal communication).

Electron Donors Anoxygenic phototrophic bacteria of the a- and b-Proteobacteria use a wide variety of reduced organic carbon compounds as electron-donating substrates (see Carbon Metabolism in this Chapter; Table 4; Fig. 8). Most phototrophic sulfur bacteria are capable of using sulfide as photosynthetic electron donor. Other inorganic electron donors utilized include H2, polysulfides, elemental sulfur, thiosulfate, sulfite, and iron (Widdel et al., 1993). Sulfide is oxidized to zero-valent sulfur, which in Chromatiaceae appears to be deposited as polysulfides or polythionates rather than in the form of S8 rings (Steudel, 1989; Steudel et al., 1990). In addition, thiosulfate is formed as an oxidation product by some species (see The Family Chlorobiaceae in Volume 7; Steudel et al., 1990). The photosynthetic sulfide oxidation rates of purple sulfur bacteria are higher than required for growth and remains constant at all growth rates. As a result, storage of sulfur is at maximum at low growth rates (van Gemerden and Mas, 1995). Zero-valent sulfur is further oxidized to sulfate. In microbial mats, polysulfides and organic sulfur compounds may be significant as photosynthetic electron donor. Polysulfide oxidation has been reported for Chlorobium limicola f.sp. thiosulfatophilum, Allochromatium vinosum, Thiocapsa roseopersicina, while dimethylsulfide is utilized and oxidized to dimethylsulfoxide by the two purple sulfur bacteria Thiocystis sp. and Thiocapsa roseopersicina (van Gemerden and Mas, 1995). In addition to reduced sulfur compounds, hydrogen serves as electron donor in the majority of green sulfur bacteria, and in the metabolically more versatile species of purple sulfur bacteria (such as Allochromatium vinosum, Thiocapsa roseopersicina). In green sulfur bacteria which lack assimilatory sulfate reduction, a reduced sulfur source is required during growth with molecular hydro-

CHAPTER 1.3

gen. Finally, a few species of purple nonsulfur bacteria, of Chromatiaceae, and of the green sulfur bacteria have been found to utilize ferrous iron as photosynthetic electron donor (Widdel et al., 1993; Heising et al., 1999). Sulfide acts as a strong poison of PSII activity in many algae and cyanobacteria. The ability of some Cyanobacteria to conduct anoxygenic photosynthesis with sulfide as an electron donor to PSI (Cohen et al., 1975; Padan, 1979; Padan and Cohen, 1982), or to continue oxygenic photosynthesis in the presence of sulfide (Cohen et al., 1986), may be one of the key traits that extend the habitat of sulfide-utilizing cyanobacteria into the temporarily anoxic, sulfide-containing, layers of hot springs (Castenholz and Utkilen, 1984), marine microbial mats (De Wit and van Gemerden, 1987a; De Wit et al., 1988), and the chemoclines of meromictic lakes (Jørgensen et al., 1979; Camacho et al., 1996). Sulfide is an inhibitor of PSII and induces the synthesis of a sulfide-oxidizing enzyme system. In contrast to phototrophic sulfur bacteria, cyanobacteria oxidize sulfide to elemental sulfur or thiosulfate but do not form sulfate (De Wit and van Gemerden, 1987b). However, the use of sulfide by cyanobacteria in anoxygenic photosynthesis must be regarded as a detoxification mechanism, since their low affinity for sulfide (De Wit and van Gemerden, 1987b; Garcia-Pichel and Castenholz, 1990) renders them unable to compete with purple or green sulfur bacteria for sulfide as an electron donor. In the natural habitat, growth of phototrophic sulfur bacteria is limited mainly by light and sulfide. Sulfide often becomes the growth-limiting factor at the top of the phototrophic sulfur bacterial layers where light intensities are highest, while sulfide has to diffuse through the remainder of the community. The affinity for sulfide during photolithotrophic growth varies between the different groups of anoxygenic phototrophs (including cyanobacteria growing with sulfide) and has been shown to be of selective value during competition experiments. Green sulfur bacteria and Ectothiorhodospiraceae exhibit 5 to 7 times higher affinities for sulfide than Chromatiaceae (van Gemerden and Mas, 1995). On the contrary, affinities for polysulfides are comparable between green sulfur bacteria and Chromatiaceae.

Efficiency of Growth and Maintenance Energy Requirements For any photochemical reaction, the quantum yield is defined as the number of molecules converted per light quantum absorbed. The quantum efficiency is the ratio of energy stored in a com-

CHAPTER 1.3

pound, to the radiant energy absorbed for its formation. The quantum requirement is the reciprocal of the quantum yield. For CO2 fixation of purple sulfur bacteria, a quantum requirement of 8 and 10.5 mol quanta· (mol CO2)-1 is theoretically expected (Brune, 1989), considering that reverse electron transport is necessary. Experimentally, a quantum requirement of 12 ± 1.5 and 11.7 mol quanta·(mol CO2-1 was determined, which corresponds to a quantum yield of 0.083 (Wassink et al., 1942 in Brune, 1989; Göbel, 1978). In contrast, calculated values for the quantum requirements of green sulfur bacteria lie between 3.5 and 4.5 mol quanta·(mol CO2)-1, if noncyclic electron transport is assumed. However, earlier measurements had yielded much higher values (9–10; Brune, 1989). This discrepancy may be explained by the very recent finding that a gene for ferredoxin-NADP+ oxidoreductase does not seem to be present in the genome of Chlorobium tepidum (D. A. Bryant, personal communication), which makes noncyclic electron transport rather unlikely also for green sulfur bacteria. The quantum yield for CO2-fixation determined for Prochlorococcus isolates incubated in daylight spectrum fluorescent light was between 0.086 and 0.128 mol C·(mol quanta)–1 (Moore et al., 1998), thus reaching Emerson’s theoretical maximum for O2 evolution in oxygenic photosynthesis. In cyanobacteria, typically thriving in oxic environments where only oxidized sources of nitrogen and sulfur are available, a large proportion of the reducing power generated in the light reactions must be diverted to assimilatory nitrate or sulfate reduction, or to nitrogen fixation, so that the quantum requirement for CO2 fixation can be substantially lower than that for oxygen evolution. In a careful study of Rhodobacter capsulatus and Rba. acidophilus grown with lactate as electron donor in a light chemostat, a value for the maintenance light energy requirement of mq = 0.012 mol quanta·(g dry weight·h)–1 was determined (Göbel, 1978). The maintenance energy requirements of green sulfur bacteria are significantly lower compared to their purple conterparts (van Gemerden and Mas, 1995). This may be explained by the fact that protein turnover is highly energy demanding and that the protein content of the green sulfur bacterial antenna is much lower than in purple sulfur bacteria (Table 3).

Response to Changes in Light Intensity and Quality Phototrophic bacteria acclimate to changes in light intensity and quality by diverse mechanisms. Anoxygenic phototrophic bacteria as well

The Phototrophic Way of Life

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as cyanobacteria respond to a step-down in irradiance by increasing the specific pigment content and vice versa (references compiled in Sánchez et al., 1998). These changes can be accomplished either by varying the number of photosynthetic units per cell, the size of the individual photosynthetic unit, or both (see Long-term Adaptations to Changes in Light Intensity in this Chapter). Besides long-term biochemical changes in the composition and the amount of light-harvesting complexes, short-term redistribution of antenna capabilities (see State Transitions in this Chapter) occur in oxygenic phototrophs. Many species use vertical migration, mediated by tactic responses (see Movement by Flagella in this Chapter) and formation of gas vesicles to regulate their vertical position and exposure to light. Especially in the stably stratified pelagic habitats of phototrophic sulfur bacteria, the difference in buoyant density from the surrounding water would cause a sedimentation of bacterial cells out of the photic zone and towards the lake bottom. The minimum buoyant density, which has been determined for phototrophic cells devoid of gas vesicles, was 1010 kg·m–3 (Overmann et al., 1991b). Actively growing cells, which contain storage carbohydrate and—in the case of Chromatiaceae—elemental sulfur, can easily attain much higher buoyant densities (up to 1046 kg·m–3; Overmann and Pfennig, 1992). By comparison, freshwater has a considerably lower density (e.g., 996 kg·m-3; Overmann et al., 1999c). As a consequence, sedimentation losses are significant for natural populations of several species of phototrophic sulfur bacteria (Mas et al., 1990). Phototrophic bacteria have developed two ways to adjust their vertical position along gradients of light intensity and spectral composition. For purple sulfur bacteria, motility in response to changes in irradiance is known to be of ecological significance in both planktonic and benthic situations. In benthic and terrestrial cyanobacteria, vertical locomotion by gliding is common. Planktonic cyanobacteria inhabiting stratified waters perform vertical migrations by changing their cellular gas vesicle content and ballast mass (intracellular carbohydrates and protein) and hence their buoyant density. Planktonic anoxygenic phototrophic bacteria do not seem to perform vertical migrations mediated by changes in gas vesicle content but rather use these cell organelles t o maintain their position within the chemocline (Overmann et al., 1991b; Overmann et al., 1994; Parkin and Brock, 1981).

Long-Term Adaptations to Changes in Light Intensity In those photosynthetic bacteria in which the entire photosynthetic apparatus is confined to

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the membrane, light absorption often is increased by formation of intracellular membrane systems (Fig. 2). In Rhodobacter capsulatus, the number of intracellular membrane vesicles increases by a factor of 6.3 when the cells are shifted from high to low light intensities. As a result, the area of intracellular membranes under these conditions is 2.7-fold larger than the area of the whole cytoplasmic membrane. Photosynthetic species of the b-Proteobacteria which do not form extensive intracellular membrane systems (Rhodocyclus purpureus, Rhodocyclus tenuis, Rubrivivax gelatinosus) increase the density of photosynthetic units in their cytoplasmic membrane (Drews and Golecki, 1995). Intracellular membranes appear to be absent in Heliobacteriaceae and Heliothrix, where pigments are confined to the cytoplasmic membrane (Fig. 2). In Chloroflexus aurantiacus, the increase in cellular concentrations of bacteriochlorophylls a and c is mediated by an increase in the number and volume of chlorosomes, and the percentage of cell membrane surface covered by chlorosomes (Golecki and Oelze, 1987). In a similar manner, green sulfur bacteria adapt to low light intensities by increasing the size and the cellular number of chlorosomes (see The Family Chlorobiaceae, Physiology section in Volume 7). During induction of the photosynthesis apparatus in Proteobacteria, invaginations of the cytoplasmic membrane, increases in the number and size of the photosynthetic units, and bacteriochlorophyll synthesis occur simultaneously. Under anoxic conditions, the amount of pigment synthesized by anoxygenic phototrophic bacteria is inversely related to the available light intensity and varies by a factor of up to 6.6 (Göbel, 1978). After a shift to low light intensity, the ratio of light-harvesting complex I per reaction center remains constant (at about 30 bacteriochlorophylls per reaction center), whereas the relative amount of the peripheral light-harvesting complex II increases. As a result, the size of the photosynthetic unit changes by a factor of two to five. Conversely, the specific NADH dehydrogenase activity decreases as does the amount of cytochrome and ubiquinone per reaction center. In Rba. capsulatus and Rba. spheroides these changes take about 2–3 generations and the growth rate is lowered during adaptation due to energy limitation. In the purple sulfur bacterium Allochromatium vinosum, low-light adaptation is also accomplished by increasing the size of the photosynthetic unit (Sánchez et al., 1998). Species like Rhodospirillum rubrum and Blastochloris viridis, which harbor only one type of light-harvesting complex, increase the number of photosynthetic units (Drews and Golecki, 1995). Similar to anoxygenic phototrophic bacteria, changes in both the number and the size of the

CHAPTER 1.3

photosynthetic unit have also been decribed for cyanobacteria. In marine Synechococcus strains, the cellular content of the light-harvesting phycoerythrin can be varied by a factor of 20 and decreases with increasing light intensity. In marine benthic Microcoleus chthonoplastes, an increase in the content of total phycobilines and a change in the ratio of PEC to PC occurs with decreasing light intensity. The latter increase the ratio of phycocyanin to chlorophyll a during low-light adaptation (Foy and Gibson, 1982; Post et al., 1985). Acclimation to very low light intensities usually involves an increase in the size of the photosynthetic unit, such as in metalimnetic Oscillatoria (Leptolyngbya) redekei and Oscillatoria agha rdii. Changes in both the number and the size of the photosynthetic units seem to occur in Microcystis (Zevenboom and Mur, 1984).

Adaptations to Low Light Intensities The capability to adapt to low light intensities represents a competitive advantage for phototrophic organisms. An estimate of the minimum irradiance Imin required for survival of phototrophic cells in the environment can be calculated from a few physiological parameters, namely the pigment content of the cells, P (in mg bacteriochlorophyll·g C–1); the maintenance energy requirement, mq (in mol quanta·g C–1·s-1); the (bacterio)chlorophyll-specific attenuation coefficient, k (in m2·mg BChl a–1); the cellular dry weight content, D (in g C·m-3); and the mean optical pathlength of one cell d: Imin = mq · D · d /[1 – exp(–k · D · P · d)] Employing the appropiate values for mq (see Efficiency of growth and maintenance energy requirements), k and P (see Light Energy and the Spectral Distribution of Radiation in this Chapter), D (1.21·105 g C·m-3; Watson et al., 1977) and d (0.5 m for the smaller anoxygenic phototrophs), this yields a minimum irradiance (Imin) of 2 mmol quanta·m–2·s–1. In many natural habitats of anoxygenic phototrophic bacteria, irradiances of this order of magnitude or lower have been measured. Prochlorococcus has been found at deep water layers down to 300 m. However, these bacteria do not grow at light intensities below 3.5 mol quanta·m–2·s-1 (Moore et al., 1998) and thus appear to be less lowlight adapted than the green sulfur bacterial strain MN1 isolated from the Black Sea which grows a t light intensities as low as 0.25 mmol quanta·m–2·s-1 (Overmann et al., 1991a). Lower irradiances could be used by phototrophic prokaryotes after a decrease of mq or an increase of P or both. Both adaptations are present in strain MN1 (Overmann et al., 1991a).

CHAPTER 1.3

Adaptations to High Light Intensities Sessile cyanobacteria living on the surface of benthic microbial mats are typically adapted to very high light conditions and contain large amounts of sunscreen pigments. For oxygenic phototrophs, special adaptations to oxygendependent photoinhibition of photosynthesis are of particular relevance. The protein D1 of PSII, coded by the psbA gene, has been identified as the central target of photoinhibition at high light intensities. In Synechococcus PCC 7942, psbA contains actually a multigene family coding for three different forms of the protein D1, which are differentially expressed according to the light conditions. Analysis of mutants showed that the isoforms expressed under high light conditions allow for optimal performance of PSII under photoinhibitory conditions (Golden, 1994). In addition, carotenoids probably play a central role in avoiding oxygen-mediated pho tosensitized bleaching of photosynthetic pigments and photooxidation of fatty acids under high light conditions.They function as antioxidant quenchers of excited molecules (such as triplet state chlorins and singlet oxygen) in many organisms and perhaps also as inhibitors of free-radical reactions (Britton, 1995). The photoprotective xanthophyll cycle typical of green algae and higher plants is not present in cyanobacteria, but judging from its increased specific content at high light intensity, zeaxanthin seems to play an important photoprotective role in some strains (Kana et al., 1988; Masamoto and Furukawa, 1997; Millie et al., 1990). Glycosylated myxoxanthophylls seem to attain the same role in others (Nonnengießer et al., 1996; Garcia-Pichel et al., 1998; Ehling-Schulz et al., 1997). Because there is a considerable photooxidation of carotenoids themselves at high light intensities, the maintenance of high carotenoid contents requires an increased expression of their biosynthetic genes.

Chromatic Adaptation Several species of cyanobacteria are capable of changing the amount of peripheral phycoerythrin in response to changes in the spectral composition of light. During growth in white or green light, red-pigmented PE hexamers are added to the peripheral rods whereas additional bluepigmented PC is added under red light (Sidler, 1994). This complementary chromatic adaptation is found only in strains capable of forming PE, but not in those forming PEC. The complementary change in antenna pigment composition optimizes the light-harvesting capabilities of populations of Oscillatoria spp., which thrive in deeper layers of stratified lakes where light is

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predominantly in the blue-green to green wavelength range (Utkilen et al., 1985; Fig. 6).

Genetic Regulation in Response to Light The synthesis of the photosystem is especially energy consuming because of the high amount of light-harvesting and reaction center protein present in phototrophically grown cells of phototrophic Proteobacteria (20% in purple nonsulfur bacteria). The maintenance energy requirements seem to be increased in low-light adapted cells (Sánchez et al., 1998). An effective regulation of photosynthesis gene expression therefore would prevent futile synthesis of cellular proteins. The synthesis of the photosystem in anoxygenic phototrophic bacteria is under the control of a complex regulatory network (Bauer and Bird, 1996). The expression of light-harvesting complex I and reaction center genes is controlled 1) by the linkage of genes in superoperons, 2) at the level of transcription initiation, and 3) posttranscriptionally by the decay rate of mRNA (Bauer, 1995). In Rhodobacter capsulatus, the genes coding the structural, biosynthetic and regulatory proteins for light-harvesting I and reaction center complexes are found assembled in a 46 kb-long photosynthetic gene cluster (Alberti et al., 1995). The arrangement of the genes within the cluster seems to be conserved among different phototrophic species of the a-Proteobacteria, like Rhodobacter sphaeroides, Rhodocista centenaria and Rhodospirillum rubrum (Bauer et al., 1993). Only the pucBA operon which codes for structural a- and b-polypeptides of light-harvesting complex II is found in a distant location on the bacterial chromosome (about 18 kb of the puhA in Rhodobacter capsulatus; Suwanto and Kaplan, 1989). In anoxygenic phototrophic bacteria, transcription of the photosynthesis genes occurs only under anoxic conditions. Different photosynthesis genes exhibit varying levels of expression and degrees of regulation (Bauer and Bird, 1996). The pufA,B,L,M genes (coding for the a- and bpolypeptide of the light-harvesting complex I and the reaction center L and M structural polypeptides) as well as puhA (coding for the structural polypeptide subunit H) are tightly coregulated, transcribed at a high rate under anoxic conditions and strongly regulated (15- to 30-fold). An inverted repeat sequence located between pufA and pufL affects the longevity of the respective mRNA primary transcript. A reduction of light leads to an activation of puf and puh gene expression by the hvrA gene product, which probably directly interacts with the two promoter regions. Light of 450 nm exhibits

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the most severe repressing effect, indicating that a flavin-binding protein (possibly HvrA itself) is the photoreceptor. Notably in aerobic phototrophic bacteria, a blue light sensitive system seems to regulate biosynthesis of bacteriochlorophyll a (Shimada, 1995). The intracellular bacteriochlorophyll concentrations appear to affect puf and puc gene expression not only at the transcriptional but also the posttranscriptional level in Rhodobacter capsulatus (Rödig et al., 1999). The polycistronic organization allows the coordinate expression of the structural polypeptides of light-harvesting complex I and the two integral membraneproteins of the reaction center. Since, however, many light-harvesting I complexes are required per reaction center in Proteobacteria, additional regulatory mechanisms must exist. Differential degradation of various portions of the polycistronic mRNA are one means to regulate the stoichiometry of different components of the photosynthetic apparatus. The synthesis of different amounts of gene products is achieved by posttranscriptional regulation (Rödig J. et al., 1999). Because of a h ighly stable secondary terminator structure at its 3´-end and the absence of specific recognition sites for endonucleolytic cleavage, the mRNA coding the two lightharvesting polypeptides has much higher stability than that of the entire puf gene transcript. The degradation of the downstream pufLM section of the mRNA is mediated by an endonuclease. A similar regulation mechanism may exist for the polycistronic mRNA of bacteriochlorophyll synthesis genes (bchFNBHLM-F1696) and the puhA, and operate in regulation of lightharvesting complex II expression. A shift to low light intensities results in an increase especially of light-harvesting complex II. The corresponding pucBA operon is highly expressed but only moderately regulated (4fold). In the purple nonsulfur bacterium Rhodobacter capsulatus, four-fold less puc mRNA but at the same time four times as many lightharvesting II complexes were detected after a shift from high to low-light conditions (Zucconi and Beatty, 1988). Therefore regulation by light most likely involves posttranscriptional regulation. A posttranscriptional regulation appears to occur (Bauer, 1995). Bacteriochlorophyll and carotenoid biosynthesis genes are only weakly expressed and moderately (2 to 4-fold) regulated. Light intensity may control the rate of bacteriochlorophyll degradation (by oxidative degradation of bacteriochlorophyll; Biel, 1986) rather than the rate of synthesis (Biel, 1995). This is another distinct difference from the regulation by oxygen, where inhibition of d-aminolevulinate synthase by molecular oxygen appears to occur (see

CHAPTER 1.3

Chemotrophic Growth with O2 in this Chapter). Bacteriochlorophyll may be stabilized by insertion in pigment-protein complexes, however. The promotor of the bacteriochlorophyll synthesis gene bchC is of the sigma-70 type and leads to one large superoperon (Yurkov and Beatty, 1998). In contrast, an alternative sigma factor appears to recognize the strongly regulated structural puf and puh genes (Bauer, 1995). These differences explain the independent and different levels of regulation observed for the two classes of genes. Recently the promoter for the carotenoid biosynthesis genes crtB and crtP were identified in Synechocystis PCC 6803, and shown to be light regulated (Fernández-González et al., 1998).

State Transitions In cyanobacteria, state transitions involve redirecting the pathways of excitation energy transfer from light-harvesting complexes to both photosystems, and can be recognized by fluorescence analysis. Cyanobacteria can reach two energetically different states, in which one of the photosystems is preferentially excited. This is achieved with fast changes in the coupling between the light-harvesting complexes and the reaction center (van Thor et al., 1999). Evidence is accumulating that at least in the chlorophyll bcontaining phototrophic bacteria (“Prochlorophytes”), the short-term regulation occurs by a mechanism similar to that in green chloroplasts (Matthijs et al., 1994). In the latter, polypeptides of the PSII antenna (LHCII) are rapidly phosphorylated during overexcitation of this photosystem, and as a consequence detach from PSII and migrate to the stromal thylakoids. This mechanisms ensures a bala nced energy distribution between PSII and PSI. The net result of state transitions is the balanced function of both photosystems and an optimization of the quantum yield for photosynthesis during short-term changes, such as those that planktonic cells might experience during vertical transport by water currents.

Movement by Flagella Phototrophic Proteobacteria swim by means of flagella, whereas one species of the green sulfur bacteria (Chloroherpeton thalassium), members of Chloroflexus subgroup and cyanobacteria move by gliding. Of the a-Proteobacteria, most phototrophic species are motile. Peritrichous or lateral flagella are only found in Rhodomicrobium vannielii and the swarming phase of Rhodocista centenaria. About two thirds of the known Chromatiaceae species are motile. Larger forms (Chromatium okenii, Chr. weissei, Chr. warmingii, Chr. buderi, Thiospirillum jenense)

CHAPTER 1.3

are motile by means of bipolar multitrichous tufts of flagella. Thiospirillum jenense is bipolarly flagellated. Forms with smaller cells are monotrichously flagel lated (small Chromatium species, Lamprocystis, Thiocystis, Thiorhodococcus, Thiorhodovibrio). All Ectothiorhodospiraceae are flagellated. A new mode of motility has been described for a unicellular cyanobacterium which moves in a similar fashion to flagellated bacteria but apparently lacks a flagellum (Waterbury et al., 1985). True phototaxis is the ability to move towards or away from the direction of light. Cyanobacteria are the only prokaryotes displaying true phototaxis (Garcia-Pichel and Castenholz, 1999). Phototaxis may not be of competitive value for microorganisms adapted to live at low light intensities in the subsurface of sediments, soils and mats because the light fields may be close to diffuse deep below the surface. However, directed movements can still be of much use in microorganisms dwelling at or close to the sediment surface, where the light fields contain a significant downward directionality. Photophobic responses are changes in the direction of movement in reaction to abrupt changes in light intensity (Castenholz, 1982; Häder, 1987). In the step-up photophobic response, organisms will reverse direction when sensing an increase in light intensit y, which results in a net accumulation of organisms at lower light intensities. In a step-down photophobic (or scotophobic) response, the organisms will tend to accumulate in the region of higher light intensity. Photophobic responses are the basis of photomovement in all flagellated bacteria (Armitage, 1997), and in most gliding cyanobacteria (Castenholz, 1982). In swimming cells of phototrophic Proteobacteria, a decrease in light intensity triggers a reversal of flagellar rotation (Rhodospirillum rubrum, Chromatium spp.) or an increase in stopping frequency (Rhodobacter sphaeroides). Owing to a memory effect, cells of the latter species retain a higher stopping frequency for up to 2 min, which prevents the cells from being trapped in the dark but instead permits reorientation of the cells and a return to higher light intensities (Armitage et al., 1995). As a result of this scotophobic response, the cells accumulate in the light and at wavelengths corresponding to the absorption maxima of photosynthetic pigments. A change in light intensity of as little as 2% can be sensed (Armitage et al., 1995). Active electron transport is required for the scotophobic response. The formation of flagella in Chromatium species is induced by low sulfide concentrations and low light intensities. These two environmental variables are mutually dependent: the lower the light intensity, the higher the sulfide concentration at which a given strain can persist in its

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motile stage (Pfennig and Trüper, 1989). In the natural environment of purple sulfur bacteria, gradients of light and sulfide are opposed to each other. The control of motility by the two interdependent environmental variables (instead of only one) enables Chromatium cells to return either from low sulfide/high light environment above the chemocline or from the high sulfide/ low light environment below the chemocline back to their habitat. In its pelagic habitat, Chromatium okenii may display diurnal migrations with a vertical amplitude of about 2 m (Sorokin, 1970). In other lakes, vertical migrations of Chromatium minus extended over a distance of 30–35 cm (Lindholm et al., 1985; Pedrós-Alió and Sala, 1990). Vertical migration of nonthermophilic Chromatium, and of Chromatium tepidum also has been observed in ponds and in intertidal or hot spring microbial mats (Castenholz and Pierson, 1995; Jørgensen, 1982; Pfennig, 1978). In the latter environments, Chromatium cells migrate upwards to the surface of the mat and enter the overlaying water as a result of positive aerotaxis during the night. The cells contain high amounts of intracellular sulfur globules, which are formed during incomplete sulfide oxidation by anoxygenic photosynthesis during daytime. It is assumed that migration into microoxic layers enables the cells to grow chemoautotrophically by oxidation of sulfide or intracellular sulfur with molecular oxygen (Jørgensen, 1982; Castenholz and Pierson, 1995). If phototrophic sulfur bacteria would solely follow the light gradient, their scotophobic response would ultimately lead them into oxic water layers. Both the scotophobic behavior and aerotaxis respond to the rate of intracellular electron flow (presumably sensed as changes in the redox state of an intermediate). Because the two tactic reponses interact through a common signal, a combination of light and molecular oxygen elicits a differential response. Rhodobacter sphaeroides exhibits pronounced aerotaxis when precultivated aerobically, but negative aerotaxis when grown anaerobically in the light. Conversely, cells only swim towards higher light intensities in anoxic medium. A pulse of oxygen in the light causes a transient fall in the membrane potential which probably represents the primary tactic signal. As a result, the bacteria move towards environments where electron transport rate is increased (Armitage et al., 1995). Rhodocista centenaria exhibits a characteristic swarming behavior. In liquid media, cells move with a single polar flagellum. Upon contact with solid agar media, formation of a large number of lateral flagella is induced. Lateral flagella allow whole colonies to swarm towards or away from the light (Ragatz et al., 1994). The supposedly

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true phototaxis of these swarming colonies (Ragatz et al., 1995) has later been proven to actually be aerotaxis following microgradients within the colonies (Sackett et al., 1997). The light sensing system in this species appears to be more complex, since infrared light leads to positive, and visible light to negative phototaxis. In microbial mats, infrared light penetrates to much greater depths than light of the visible wavelength range (see Competition for Light in this Chapter). It has been suggested that the ratio of visible to in frared light may be used to maintain an optimum position in such environments (Armitage et al., 1995; Ragatz et al., 1995). Cyanobacteria are the only prokaryotes displaying true phototaxis (Garcia-Pichel and Castenholz, 1999). Surface dwelling cyanobacteria such as Lyngbya spp. from hot springs mats and intertidal sediments and the motile phases (hormogonia) of terrestrial Nostoc spp. from desert soils exhibit this type of movement. The bundleforming Microcoleus chthonoplastes also is able to display a “populational phototaxis” in that bundles of trichomes of this cyanobacterium are able to steer in the direction of the incoming light, whereas single trichomes are apparently not able to do so (Prufert-Bebout and GarciaPichel, 1994). True phototaxis is a mechanism for the orientation of cells at or close to the sediment surface, where the light field contains a significant downward directionality. In contrast, phototaxis does not provide a selective advantage for bacteria thriving in the subsurface of sediments, soils and mats because of the diffuse light field. In natural microbial mats photophobic responses to changes in light intensity are probably involved in the migrations of gliding bacteria (Nelson and Castenholz, 1982; Pentecost, 1984). In microbial mats, some strains of cyanobacteria are able to migrate vertically following their optimal light intensity over the diel cycle (Garcia-Pichel et al., 1996). The upward migrations of cyanobacteria in mats is preferentially prevented by short wavelengths, especially by UV radiation (Garcia-Pichel and Castenholz, 1994b; Bebout and Garcia-Pichel, 1995; Kruschel and Castenholz, 1988) and not by red nor green light. Phototrophic consortia are structural associations between a colorless central bacterium and several surrounding cells of pigmented epibionts (see Interactions between Phototrophic Bacteria and Chemotrophic Bacteria in this Chapter; The Family Chlorobiaceae in Volume 7; Fig. 5). Intact consortia of the type “Chlorochromatium aggregatum” exhibit a scotophobic response and accumulate in a spot of white light. In phototrophic consortia, only the central colorless bacterium carries a flagellum (J. Glaeser and J. Overmann, unpublished observation). The action spectrum

CHAPTER 1.3

of scotophobic accumulation corresponds to the absorption spectrum of the green sulfur bacterial epibionts, however. It has to be concluded that a rapid signal transfer exists between the lightsensing but immotile epibionts and the colorless motile rod (Fröstl and Overmann, 1998).

Gas Vesicles Buoyancy-conferring gas vesicles are common in green sulfur bacteria, Chromatiaceae, and cyanobacteria. Gas vesicles are cylindrical structures with conical ends; their length and width are variable and species-specific. The sheath of gas vesicles are composed of proteins (Walsby, 1994). The gas mixture within the gas vesicles is the same as in the surrounding medium and is at the same partial pressures. Gas vesicles occur in a third of the species of Chromatiaceae (belonging to the genera Amoebobacter, Lamprobacter, Lamprocystis, Thiodictyon, Thiopedia, Thiolamprovum) and some green sulfur bacteria (genera Ancalochloris, Pelodictyon, Chloroherpeton). Of the Ectothiorhodospiraceae, only Ectothiorhodospira vacuolata forms gas vesicles during stationary phase. This reflects the distribution of both families of purple sulfur bacteria in nature, where Chromatiaceae typically colonize lowlight stratified aquatic environments, whereas Ectothiorhodospiraceae typically inhabit more shallow saline ponds and sediments. Gas vesicles also are present in Prochlorothrix hollandica. In planktonic habitats, cells of cyanobacteria and phototrophic sulfur bacteria often contain gas vesicles, which indicates a selective advantage of this cellular property. Gas vesicle formation in the green sulfur bacterium Pelodictyon phaeoclathratiforme is induced exclusively at light intensities 100 s-1 in Rhodobacter sphaeroides and requires between 200 and 1000 H+ per rotation (Armitage et al., 1995). This yields a proton translocation rate of ~6 ¥ 104 H+·s-1 at a swimming velocity of 100 m·s-1. Based on an absorbing cross sectional area of the cell of 1 m2, an absorption of 36% of the incident light (see Efficiency of Light Harvesting in this Chapter), a ratio of protons translocated to electrons transferred (H+/e- ratio) of 2 (see Conversion of Light into Chemical Energy in this Chapter), and assuming that each photon absorbed leads to transport of an electron, the proton translocation rate of 6 ¥ 104 H+·s-1 would be reached at an underwater irradiance of 0.2 mol quanta·m-2·s-1. However, all available quanta would be required just for motility at this irradiance and no vertical migration would be possible during the night. Therefore motility by flagella will be of compet-

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itive advantage only at significantly higher irradiances. In many lakes, underwater irradiances in layers of phototrophic sulfur bacteria are £1 mol quanta·m–2·s-1 (Overmann and Tilzer, 1989a; Overmann et al., 1999a). Under these conditions, purple sulfur bacteria harboring gas vesicles dominate over flagellated forms in the chemocline community (Fig. 4). At least in some lakes, gas vesicles appear to be of selective advantage also at higher underwater irradiances (Overmann et al., 1991b; Overmann and Pfennig, 1992). Interestingly, the extremely low-light adapted Chlorobium phaeobacteroides strain MN1 isolated from the chemocline of the Black Sea was not capable of gas vesicle formation. The green sulfur bacterial layer is located at an 80-m depth and with respect to light intensity represents the lower limit for growth of a phototrophic organism (see The Family Chlorobiaceae in Volume 7). The isolated strain exhibits an extremely low maintenance energy requirement. It therefore appears that gas vesicle formation is too energy demanding at the very low light intensities available at an 80-m depth in the Black Sea.

Carbon Metabolism of Phototrophic Prokaryotes In the natural environment, the principal carbon source of phototrophic bacteria in many instances is CO2 (Madigan et al., 1989; Sinninghe Damsté et al., 1993; Takahashi et al., 1990). In Cyanobacteria, Chromatiaceae, Ectothiorhodospiraceae and purple nonsulfur bacteria, CO2 is assimilated by the reductive pentose phosphate or Calvin cycle. Employing this cycle, the formation of one molecule of glyceraldehyde-3phosphate requires 6 NAD(P)H+H+ and 9 ATP. By comparison, the reductive tricarboxylic acid cycle used for CO2-assimilation by green sulfur bacteria requires 4 NADH+H+, 2 reduced ferredoxins, and only 5 ATP. As two of the reactions of the reductive tricarboxylic acid cycle (the a-oxoglutarate synthase and pyruvate synthase rea ctions) require reduced ferredoxin as electron donor, this pathway of CO2 fixation can only proceed under strongly reducing conditions. Furthermore, reduced ferredoxin is a primary product of the light reaction only in FeS-type reaction centers. Ultimately, the lower demand for ATP is possible because of the adapatation of green sulfur bacteria to the strongly reducing conditions of their natural environment. CO2fixation by the hydroxypropionate cycle in Chloroflexus aurantiacus requires 8 ATP per glyceraldehyde-3-phosphate and therefore is energetically less favorable than in green sulfur bacteria.

CHAPTER 1.3

Organic carbon as it is present in canonical microbial biomass (; Harder and van Dijken, 1976) is considerably more reduced than CO2. Given the high energy demand of autotrophic growth, the capability for assimilation of organic carbon compounds is of selective advantage especially if natural populations are limited by light or by low concentrations of electron-donating substrates, as is typically the case for phototrophic sulfur bacteria. At limiting concentrations of sulfide or thiosulfate, the cell yield of green sulfur bacteria is increased three times if acetate is available as an additional carbon source (Overmann and Pfennig, 1989b). Acetate represents one of the most important intermediates of anaerobic degradation of organic matter (Wu et al., 1997). That almost all anoxygenic phototrophic bacteria (with the exception of Rhodopila globiformis; Imhoff and Trüper, 1989) are capable of acetate assimilation is therefore not surprising. In most phototrophic Proteobacteria, acetate is assimilated by acetylCoA synthetase and the enzymes of the glyoxylate cycle. In green sulfur bacteria, the ferredoxin-dependent pyruvate synthetase, PEP synthetase, and reactions of the reductive tricarboxylic acid cycle serve this purpose. The capacity for organotrophic growth seems to correlate with the presence of a-oxoglutarate dehydrogenase. The latter is a key enzyme for the complete oxidation of the carbon substrates in the tricarboxylic acid cycle (Kondratieva, 1979), whereas a complete cycle is not needed for the photoassimilation during the presence of inorganic electron donors. The range of carbon substrates utilized and the capacity for photoorganotrophy or chemoorganotrophy varies considerably among the different groups of phototrophic pr okaryotes (Pfennig and Trüper, 1989). Organic carbon compounds not only are assimilated but also can serve as photosynthetic electron donors in purple nonsulfur bacteria, some Chromatiaceae and Ectothiorhodospiraceae, all Heliobacteriaceae, and members of the Chloroflexus subdivision. Green sulfur bacteria are the least versatile of all phototrophic prokaryotes. All known species are obligately photolithotrophic and assimilate only very few simple organic carbon compounds (acetate, propionate, pyruvate). Few strains have been shown to assimilate fructose or glutamate. Whereas green sulfur bacteria have a higher growth affinity for sulfide than purple sulfur bacteria, acetate seems to be used by purple sulfur bacteria at an affinity 30 times higher than in green sulfur bacteria (Veldhuis and van Gemerden, 1986). In addition, uptake of acetate in Chlorobium phaeobacteroides is inhibited by light (Hofman et al., 1985).

CHAPTER 1.3

Based on their metabolic flexibility, two groups can be distinguished among the Chromatiaceae. Several species (Chromatium okenii, Chr. weissii, Chr. warmingii, Chr. buderi, Chr. tepidum, Thiospirillum jenense, Lamprocystis roseopersicina, Thiodictyon elegans, Thiodictyon bacillosum, Thiocapsa pfennigii, Thiopedia rosea) are obligately phototrophic, strictly anaerobic and photoassimilate acetate and pyruvate only in the presence of CO2 and sulfide. Assimilatory sulfate reduction is absent in these species (Pfennig and Trüper, 1989). However, particularly those species with limited metabolic flexibility form dense blooms under natural conditions (see Coexistence of Phototrophic Sulfur Bacteria in this Chapter). The second physiological group within the Chromatiaceae comprises the small Chromatium species (Chr. gracile, Chr. minus, Chr. minutissimum), Allochromatium vinosum, Lamprobacter modestohalophilus, as well as Thiocystis spp., Thiocapsa. Most of these species use thiosulfate as electron donor and a wide range of organic carbon compounds including glucose, fructose, glycerol, fumarate, malate, succinate, formate, propionate, and butyrate for photoassimilation, and often are capable of assimilatory sulfate reduction. In some species (especially Allochromatium vinosum), these organic carbon substrates also serve as electron-donor for phototrophic or chemotrophic growth. Most Ectothiorhodospiraceae species are capable of photoorganotrophic growth, with Ectothiorhodospira halophila and Ectothiorhodospira halochloris being the exceptions. The spectrum of electron-donating carbon substrates for photoorganotrophic growth resembles that found in the versatile Chromatium species (Pfennig and Trüper, 1989). Assimilation of acetate and propionate proceeds by carboxylation and therefore depends on the presence of CO2. Chloroflexus aurantiacus grows preferably by photoorganoheterotrophy (Pierson and Castenholz, 1995). The carbon substrates utilized comprise acetate, pyruvate, lactate, butyrate, C4-dicarboxylic acids, some alcohols, sugars and amino acids (glutamate, aspartate). This versatility has been seen as the major cause for the profuse growth of Chloroflexus in microbial mats where accompanying microorganisms, especially cyanobacteria, may provide the required carbon substrates (Sirevåg, 1995). However, high rates of formation of low-molecular-weight organic carbon substrates by the anaerobic food chain have also been observed in other stratified systems, where the dominating anoxygenic phototrophs could utilize only a narrow range of carbon substrates (Overmann, 1997; Overmann et al., 1996). The refore, the presence of lowmolecular-weight organic carbon substrates is

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not necessarily the most selective factor in the natural environment. Slow photolithoautotrophic growth with H2S or H2 as electron-donating substrates has been shown in laboratory cultures of Chloroflexus aurantiacus and in hot spring populations (Pierson and Castenholz, 1995). Carbon fixation proceeds by carboxylation of acetyl-CoA and via hydroxypropionyl-CoA as an intermediate and yields glyoxylate as the net product (hydroxypropionate cycle; Holo, 1989; Strauß and Fuchs, 1993; Eisenreich et al., 1993). So far this cycle has not been found in any other member of the Bacteria. Glyoxylate is further assimilated into cell material with tartronate semialdehyde and 3phosphoglycerate as intermediates (Menendez et al., 1999). The highest metabolic versatility is found in phototrophic a- and b-Proteobacteria (purple nonsulfur bacteria). All representatives grow photoorganoheterotrophically and (with the exception of Blastochloris viridis) photolithoautotrophically with H2 in the light. In addition to the substrates used by versatile purple sulfur bacteria, the spectrum of substrates that can serve as electron donors comprise long-chain fatty acids (like pelargonate), amino acids (aspartate, arginine, glutamate), sugar alcohols (sorbitol, mannitol), or aromatic compounds (benzoate; Imhoff and Trüper, 1989). With the exception of Rubrivivax gelatinosus, none of the purple nonsulfur bacteria is capable of degradation of polymers and therefore depends on the anaerobic food chain for the supply of electron-donating substrates required for growth. This dependence and the competition with chemotrophs for the carbon substr ates might be the major reason why dense blooms of purple nonsulfur bacteria do not occur under natural conditions (see Habitats of Phototrophic Prokaryotes in this Chapter). Some species are capable of also using reduced sulfur compounds as electron donors. However, most species oxidize sulfide to elemental sulfur only (Hansen and van Gemerden, 1972). In Heliobacteriaceae, only a limited number of carbon substrates can serve as photosynthetic electron donor including pyruvate, ethanol, lactate, acetate, and butyrate. High levels of sulfide are inhibitory (Madigan, 1992; Madigan and Ormerod, 1995). Cyanobacteria are obligate autotrophs par excellence; however, small molecular weight organic compounds such as acetate, sugars and amino acids are assimilated. In the case of amino acids, the presence of various efficient uptake systems has been interpreted as a means of recovery of leaked organic nitrogen, rather than a true chemotrophic capability (Montesinos et al., 1997). Certain strains of cyanobacteria can grow facultatively as chemoheterotrophs in the

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dark (Rippka et al., 1979), but even under these conditions all of the photosynthetic machinery is synthesized. This lack of regulation implies that chemotrophy has played no significant evolutionary role in these organisms.

Chemotrophic Growth with O2 Ecophysiology of Chemotrophic Growth In lakes, purple sulfur and green sulfur bacteria are confined to environments where light reaches sulfide-containing water layers. The physiological properties restrict the distribution of these bacteria in the pelagic habitat (Pfennig, 1978). Dense accumulations of anoxygenic phototrophic bacteria, which apparently are growing chemotrophically, are only known for Chloroflexus (see Habitats of Phototrophic Prokaryotes in this Chapter). Although populations of other anoxygenic phototrophic bacteria do not seem to grow permanently by chemotrophy, the ability of many strains to shift to an aerobic chemotrophic mode of growth is of selective advantage in environments like intertidal sediments. Green sulfur bacteria and Heliobacteriaceae are obligate anaerobes. Under oxic conditions, the reaction of reduced ferredoxin of the type I reaction center with molecular oxygen would create superoxide and other activated oxygen species. Heliobacteriaceae are rapidly damaged by exposure to molecular oxygen. This has been attributed not only to the formation of toxic oxygen radicals but also the destruction of the unsaturated fatty acids present in the cell membrane by activated oxygen species (Madigan and Ormerod, 1995). In green sulfur bacteria, it has been observed that the energy transfer from light-harvesting bacteriochlorophylls c/d/e to bacteriochlorophyll a drops by a factor of 10 after an increase in redox potential due to the quenching by chlorobium quinone. This mechanism may protect the cells during brief anoxic/ oxic transitions. (see The Family Chlorobiaceae, Physiology section in Volume 7). All other groups of phototrophic prokaryotes comprise species that not only generate metabolic energy by photosynthesis but are also capable of chemosynthesis with O2. Chloroflexus aurantiacus is capable of growth as an aerobic heterotroph. During phototrophic growth, b-carotene, g-carotene, and hydroxy-gcarotene-glucoside are the major carotenoids, whereas echinenone and myxobactone predominate in aerobically grown cells (Pierson and Castenholz, 1995). Unlike in purple nonsulfur or purple sulfur bacteria, synthesis of some carotenoids by C. aurantiacus is greatly enhanced under aerobic conditions (Pierson and Casten-

CHAPTER 1.3

holz, 1974). The expression of the chlorosome CsmA protein is transcriptionally or posttranscriptionally regulated by oxygen (Theroux et al., 1990). Almost all known species of phototrophic aand b-Proteobacteria (purple nonsulfur bacteria) are capable of microaerophilic or aerobic chemoorganoheterotrophic growth with oxygen as terminal electron acceptor. Of the purple sulfur bacteria, Ectothiorhodospira species, and eight small-celled species of the Chromatiaceae (Thiocapsa rosea; Chromatium gracile; Chr. minus; Allochromatium vinosum; Thiocystis violascens; Thiocapsa roseopersicina; Thiocystis violacea; Thiorhodovibrio winogradskyi) can grow by chemolithotrophy, oxidizing sulfide or thiosulfate with molecular oxygen (De Wit and van Gemerden, 1987b; Kämpf and Pfennig, 1980; Overmann and Pfennig, 1992). Only few species grow also chemoorganotrophically with organic carbon substrates as electron donor of respiration. The group of facultatively chemotrophic Chromatiaceae includes typical inhabitants of benthic microbial mats like Thiocapsa roseopersicina and Thiorhodovibrio winogradskyi. This is not surprising considering the pronounced oxic/ anoxic fluctuations in this type of habitat. The cells of purple sulfur bacteria in benthic systems are often immotile and form aggregates together with sand grains, apparently as an adaptation to the hydrodynamic instability of the habitat (van den Ende et al., 1996). At the same time, however, immotile cells are exposed to strong diurnal variations in oxygen concentrations. The growth affinities for sulfide are lower for chemotrophically growing Thiocapsa roseopersicina than for colorless sulfur bacteria, which may explain that no natural populations of purple sulfur bacteria are known that grow permanently by chemotrophy (see Interactions between Phototrophic Sulfur Bacteria and Chemotrophic Bacteria in this Chapter). When grown anaerobically in the light, facultatively chemotrophic species of the purple nonsulfur and purple sulfur bacteria contain a potentially active repiratory system and exhibit ≥50% of the respiratory activity of chemotrophically growing cells (De Wit and van Gemerden, 1987a; Kämpf and Pfennig, 1980; Overmann and Pfennig, 1992; Pfennig, 1978). In cells that still contain bacteriochlorophyll, respiration is inhibited by light. This indicates that respiration and photosynthesis are coupled (e.g., by the membrane potential or common redox carriers; Richaud et al., 1986). An example is the soluble cytochrome c2 which has a dual function in Rhodobacter sphaeroides where it is needed for electron transfer from the cytochrome bc1 complex to the reaction center during ph otosynthesis, and to the cytochrome c oxidase during

CHAPTER 1.3

respiration with molecular oxygen. During photosynthetic growth, expression of cytochrome c2 is increased. At limiting concentrations of electron donating substrate, photosynthesis is preferred over respiration as long as the intracellular bacteriochlorophyll content is maintained at a sufficiently high level (4–7 g bacteriochlorophyll a·mg protein–1 in Thiocapsa roseopersicina at light saturation; De Wit and van Gemerden, 1990a). Growth continues after a shift to microoxic or aerobic conditions. Under oxic conditions the synthesis of pigments and of pigment-binding proteins of the photosynthetic apparatus ceases. The number of intracellular membrane vesicles is reduced dramatically and the composition of membrane lipids is altered. The pigment content in purple sulfur bacteria is inversely related to the ambient oxygen concentration (Kämpf and Pfennig, 1986). At 25% air saturation (52 M) of oxygen, pigment synthesis in Thiocapsa roseopersicina is completely repressed and cells become colorless (De Wit and van Gemerden, 1987b). In continuous cultures of purple sulfur bacteria, active degradation has not been observed and intracellular bacteriochlorophyll concentrations follow the washout curve. Thus bacteriochlorophyll does not seem to be actively degraded but is diluted out by cell division (De Wit and van Gemerden, 1987b). Concomitantly, the activities of respiratory enzymes (NADH dehydrogenase, cytochrome c oxidases) are increased in chemotrophically grown cells. When the cells of Thiocapsa roseopersicina become colorless, they use only one third of the electron donor for reduction of CO2. The remaining two thirds are used for energy generation and respired. Correspondingly, the protein yield reaches one third of that of phototrophically grown cells (De Wit and van Gemerden, 1987b; De Wit and van Gemerden, 1990b). In aerobic phototrophic bacteria, aerobic growth is stimulated by light that is absorbed by bacteriochlorophyll a. This stimulation is only transient, however, since bacteriochlorophyll synthesis is repressed even by low light intensities (Yurkov and van Gemerden, 1993) thus leading to a loss of the photosynthetic apparatus under continuous illumination. Respiration in cyanobacteria involves a full respiratory chain including a cytochrome aa3 terminal oxidase. Monomeric sugars are degraded using the oxidative pentose phosphate cycle. A complete tricarboxylic acid cycle has never been shown for any cyanobacterium. The NADPH formed in sugar catabolism is fed to the membrane-bound electron transport chain at the level of plastoquinone. This is in contrast to green chloroplasts, in which plastoquinol is autoxidized (Peltier and Schmidt, 1991). The res-

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piratory electron transport chain of cyanobacteria is located in both the plasma and the thylakoidal membrane, and it shares many functional components with photosynthetic electron transport. The role of exogenous respiration of organic substrates is probably minor under natural conditions. Under anoxia, the known electron acceptor alternatives to oxygen for cyanobacterial chemoorganotrophy are some organic compounds and elemental sulfur. Fermentation seems to be a relatively widespread ability in benthic and bloom-forming cyanobacteria, but it is not universal (Moezelaar and Stal, 1994).

Genetic Regulation by O2 A shift from anoxic to oxic growth conditions requires the expression of new proteins and cofactors. On the genetic level the formation of the photosynthetic apparatus and the intracytoplasmic membrane system is regulated by two main environmental variables, light intensity (see Response to Changes in Light Intensity and Quality in this Chapter) and molecular oxygen. The two factors act independently of one another and are involved in different mechanism of regulation of bacteriochlorophyll synthesis (Arnheim and Oelze, 1983). Compared to light, molecular oxygen acts as a stronger repressor, however. Although oxygen is a major factor controlling the formation of the photosynthetic apparatus in most of the facultatively phototrophic Proteobacteria, Rhodovulum sulfidophilum and Rhodocista centenaria are exceptional in that these species form the photosynthetic apparatus under both aerobic and anaerobic conditions (Hansen and Veldkamp, 1973; Nickens et al., 1996). Photopigment synthesis is not repressed by O2 in Rhodocista centenaria. The regulation of bacteriochlorophyll synthesis in purple nonsulfur bacteria is complex. The cells synthesize very little bacteriochlorophyll, probably because of the inhibition of bacteriochlorophyll biosynthesis enzymes (the d–aminolevulinic acid synthesis and enzymes for the conversion of coproporphyrin; Oelze, 1992) by O2. Oxygen does not seem to exert an effective transcriptional control. Under oxic conditions the transcription of bacteriochlorophyll synthesis genes decreases 2-fold, while that of lightharvesting I and reaction-center genes decreases by a factor of 30–100 (Bauer, 1995). The tetrapyrrole synthesis pathway has four different branches (leading to heme, bacteriochlorophyll, siroheme and vitamin B12). While the bacteriochlorophyll content is drastically reduced in the presence of oxygen (Arnheim and Oelze, 1983), heme synthesis remains unaffect ed (Lascelles, 1978). The intracellular activity of d-aminolevulinic acid synthase, the key enzyme of tetrapy-

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rrol synthesis in a-Proteobacteria, is reduced in the presence of oxygen. Regulation by oxygen may occur also during some later steps of tetrapyrrole synthesis. It appears that oxygen inhibits magnesium chelatase, thereby increasing the protoporphyrin IX pool, which in turn leads to increased formation of heme. Feedback inhibition of d-aminolevulinate synthase by heme would then slow down the synthesis of intermediates but still guarantee the amount needed for heme biosynthesis (Beale, 1995; Biel, 1995; Rebeiz and Lascelles, 1982). After return to anoxic conditions the synthesis of the photosynthetic apparatus and intracellular membranes occurs in a light-independent manner. Anoxygenic photosynthetic bacteria contain a distinct light-independent protochlorophyllide reductase, composed of probably three subunits (BchN, BchB, and BchL). In angiosperms, the reduction of the fourth ring of the Mg-tetrapyrrole intermediate by NADPHprotochlorophyllide oxidoreductase is a lightdependent step in the chlorophyll biosynthetic pathway. This protein represents one of the only two enzymatic transformations known to require light (Suzuki and Bauer, 1995). Cyanobacteria, green algae and gymnosperms contain both, the light-dependent and light-independent protochlorophyllide reductase. The capacity to synthesize (bacterio)chlorophyll in the dark is of significance for the competitive success of Chromatiaceae in intertidal microbial mats. During anoxic conditions in the dark, Thiocapsa roseopersicina can synthesize bacteriochlorophyll a at maximum rate. Under the fluctuating conditions as they are observed in benthic microbial mats (oxic light, anoxic dark phase), purple sulfur bacteria therefore can maintain a photosynthetic mode of growth as long as bacteriochlorophyll synthesis during the night compensates for the wash out of pigments during the day (De Wit and van Gemerden, 1990b). A multicomponent regulatory cascade controls the coordinate expression of the lightharvesting and reaction center puf, puh, and puc genes and involve various transcription factors (Bauer, 1995; Bauer and Bird, 1996). In Rhodobacter capsulatus, a redox-sensitive repressor (CrtJ) binds under oxic conditions to a conserved palindrome sequence in promotors of bacteriochlorophyll, carotenoid, and lightharvesting complex II genes. A second system for the regulation of the puf, puh, and puc operons probably consists of three components, a membrane-spanning sensor kinase (RegB), a soluble response regulator (RegA), and a hypothetical activator of the nonspecific alternative sigma factor sP (RegX). A decrease in oxygen tension causes autophos phorylation of the membranespanning sensor kinase RegB, which then phos-

CHAPTER 1.3

phorylates the cytoplasmic response regulator RegA. The latter acts as intermediate and probably transfers its phosphate to a putative third DNA-binding component that activates gene expression. The RegA-RegB system also is involved in regulation of the expression of cytochrome c2 and the Calvin cycle CO2 fixation genes and therefore is of general significance for the regulation of cellular metabolism. The transcripts of the photosynthetic gene cluster exceed 10 kb and extend from pigment biosynthesis genes across promoter regions and into the genes for light-harvesting complex I and reaction center proteins. In Rhodobacter capsulatus, transcription of the genes coding structural polypeptides of the reaction center and lightharvesting complex I are not the only peptides initiated at their respective promotors. The transcripts of the bacteriochlorophyll biosynthesis bchCA operon extends through the promoter and coding sequences of the downstream puf BALM operon, and the transcript of the carotenoid biosynthesis crtEF operon extends through both (Wellington et al., 1992). Similarly, the bchFBKHLM-F1696 and puhA operons are transcriptionally linked. The linkage of operons of different components of the phot osynthetic apparatus in such superoperons also has been detected in other species of purple nonsulfur bacteria and may play a significant role in the adaptation of cells to changes in environmental oxygen tension. According to a model (Wellington et al., 1992), the presence of superoperons ensures a rapid physiological response to a decrease in oxygen tension. In the presence of oxygen, a basal level of light-harvesting I and reaction center polypeptides is constantly formed and incorporated into the membrane, but these polypeptides disappear again in the absence of bacteriochlorophyll (Dierstein, 1984; Drews and Golecki, 1995) due to degradation. After a shift from oxic to anoxic conditions, the presence of a basal level of structural polypeptides considerably shortens the lag time for the change from aerobic respiratory to anaerobic photosynthetic growth. During this lag ph ase, the cellular amount of structural polypeptides of the photosynthetic apparatus is further increased by increasing the transcription rate of the puf and puh genes. Oxygen does not only regulate the transcription of photosynthesis genes but also later steps in gene expression. Posttranscriptional regulation involves mRNA processing (mRNA degradation) and possibly some later steps (Rödig J. et al., 1999). In most bacteria, the formation of multiple sigma factors is a prerequisite for the coordination of the regulation of a large number of genes in response to changes in environmental condi-

CHAPTER 1.3

tions. Sigma factors are dissociable subunits that confer promoter specificity on eubacterial core RNA polymerase and are required for transcription initiation. In phototrophic bacteria, the diversity of sigma factors of the s70 family as they are present in the different phylogenetic groups appears to be correlated with their metabolic flexibility. In the unicellular cyanobacteria Synechococcus sp. and Synechocystis sp., nine different sigma factors (one member of group 1, four members of group 2, and four members of group 3) have been found, whereas one group 1 and three group 2 sigma factors have been found in Chloroflexus spp. In contrast to most other bacteria, the green sulfur bacterium Chlorobium tepidum contains only one group 1, but no alternative group 2 sigma factor (Gruber and Bryant, 1998). In Chloroflexus, one group 2 s70 factor (SigB) is transcribed at fourfold higher levels during aerobic growth and therefore appears to be involved in the shift in metabolism. It has been proposed that SigB is involved in regulation of pigment synthesis (Gruber and Bryant, 1998).

Significance of Anoxygenic Photosynthesis for the Pelagic Carbon and Sulfur Cycles The carbon fixation of phototrophic sulfur bacteria has been determined in a wide range of habitats, mostly inland lakes (Overmann, 1997; van Gemerden and Mas, 1995). The theoretical maximum of primary production by phototrophic sulfur bacteria has been estimated to be 10,000 mg C·m–2·d–1. Purple and green sulfur bacteria can contribute up to 83% of total primary productivity in these environments. This high number notwithstanding, anoxygenic primary production only represents a net input of organic carbon to the food web if 1) the anaerobic food chain is fueled by additional allochthonous carbon from outside and 2) aerobic grazers have access to the biomass of phototrophic sulfur bacteria. Based on recent experimental evidence, these conditions are met at least in some aquatic ecosystems (Overmann, 1997). With the exception of geothermal springs, the sulfide required by phototrophic sulfur bacteria for CO2-assimilation originates from sulfate or sulfur reduction during the terminal degradation of organic matter. This organic matter cannot be provided solely by anoxygenic phototrophic bacteria, since growth (hence accumulation of reduced carbon) constantly diverts electrons from their cycling between anoxygenic phototrophic bacteria and sulfate-reducing bacteria. At least part of the sulfide formation is therefore fueled by carbon that has already been fixed by oxygenic photosynthetic organisms within or outside the ecosystem. Consequently anoxygenic photosynthesis represents not new, but second-

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ary primary production. A complete degradation of the carbon fixed by phototrophic sulfur bacteria in the anaerobic food chain (and thus an efficient recycling of electrons) in an anoxygenic primary production has been estimated to exceed oxygenic photosynthesis by as much as ten times (Overmann, 1997). In reality, anoxygenic photosynthesis surpasses that of phytoplankton mostly in oligotrophic lakes. In many oligotrophic lakes, the input of allochthonous carbon derived from terrestrial sources in the watershed is significant (Rau, 1980; Sorokin, 1970). In an oligotrophic saline meromictic lake (Mahoney Lake, B.C., Canada), purple sulfur bacteria together with the anaerobic food chain efficiently converted allochthonous organic carbon into easily degradable bacterial biomass (Overmann, 1997). It appears likely that phototrophic sulfur bacteria have this ecological function also in other aquatic ecosystems. The presence of hydrogen sulfide in layers of phototrophic sulfur bacteria may prevent their biomass from entering the grazing food chain. This has been substantiated by stable carbon and sulfur isotope data, which indicated that phototrophic sulfur bacteria are not consumed to a significant extent by higher organisms (Fry, 1986). In addition, a quantitative analysis of loss processes conducted in a few lakes indicates that predation must be of minor significance (Mas et al., 1990; van Gemerden and Mas, 1995). In contrast, recent investigations have revealed that at least in one lake ecosystem, a major fraction of purple sulfur bacterial biomass enters the aerobic food chain via rotifers and calanoid copepods (Overmann et al., 1999b; Overmann et al., 1999c). The key environmental factors that caused this efficient link between anoxic and oxic water layers were the autumnal upwelling of phototrophic bacteria into oxic water layers by mixing currents, and the formation of gas vesicles and large cell aggregates by the dominant species, Amoebobacter purpureus. Sulfide formation by sulfate- and sulfurreducing bacteria and sulfide oxidation back to sulfur and sulfate occur at comparable rates in several lakes (Overmann et al., 1996; Parkin and Brock, 1981). This leads to a closed sulfur cycle and a detoxification of sulfide without concomitant depletion of oxygen (Pfennig, 1978). The significance of phototrophic sulfur bacteria for the oxidation of sulfide in stratified environments is critically dependent on their cell density rather than the absolute biomass per surface area of the ecosystem (Jørgensen, 1982). Dense populations in laminated microbial mats can account for 100% of the total sulfide oxidation in those systems, whereas some dilute pelagic populations oxidize only very small amounts (e.g., 4% in the Black Sea) of the sulfide

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diffusing from below into the chemocline (Overmann et al., 1991a; Overmann et al., 1996). No information on the ecological significance of aerobic phototrophic bacteria is available to date.

Interactions with Other Microorganisms Competition for Light Blue light prevails in very clear open oceans (Fig. 6) where marine Synechococcus cells thrive under conditions of low photon flux (~10 mol quanta·m–2·s–1; Carr and Mann, 1994). Two ecotypes of the marine Synechococcus exist which differ in the intracellular ratio of phycourobilin to phycoerythrobilin (Waterbury et al., 1986). Two subpopulations are distinguished according to the predominant chromophore associated with the phycoerythrin. Phycourobilin-rich strains are characteristic of the open oceans whereas strains with a lower PUB content predominate in shelf waters (Olson et al., 1990a). Compared to PEB-containing antennae (absorption maximum, ~550 nm), incorporation of PUB (absorption maximum, ~495 nm) increases the efficiency of light absorption significantly in deeper water lay ers of oligotrophic oceans. Similarly, coexisting and phylogenetically closely related but genetically distinct populations of Prochlorococcus are adapted for growth at different light intensities, which results in their broad depth distribution (Moore et al., 1998). The low-light-adapted ecotype has a higher intracellular content of chlorophylls a and b, a higher chlorophyll b/a ratio, and exhibits a higher maximum quantum yield reaching the theoretical maximum of 0.125 mol C·(mol quanta)–1. Its properties enable this ecotype to colonize very low water layers. It has been suggested that the distribution of different ecotypes in the same water column would result in greater integrated production than could be achieved by a single ecotype (Moore et al., 1998). Based on the specific physiological properties of oxygenic and anoxygenic phototrophic bacteria, multilayered microbial communities frequently develop in stratified pelagic and in benthic (Fig. 5A,B) habitats. Cyanobacteria, eukaryotic algae and even plants (Lemna) form the topmost layers overlying populations of Chromatiaceae and green sulfur bacteria (Dubinina and Gorlenko, 1975; Caldwell and Tiedje, 1975; Pfennig, 1978; Camacho et al., 1996; Pierson et al., 1990; Pierson et al., 1990). Phototrophic sulfur bacteria require the simultaneous presence of light and sulfide, which usually restricts their occurrence to layers well below the surface of lakes and sediments. As a consequence of the absorption of light in the overlying water, the light energy available to phototrophic

CHAPTER 1.3

sulfur bacteria in most pelagic environments is rather low (0.02–10% of surface light intensity; van Gemerden and Mas, 1995; Parkin and Brock, 1980b; Camacho et al., 1996). Similar values have been determined for purple layers in benthic microbial mats (Kühl and Jørgensen, 1992; Pierson et al., 1990; Garcia-Pichel et al., 1994c). A tight correlation between anoxygenic photosynthesis and the amount of light reaching phototrophic sulfur bacteria strongly suggests that light i s the main environmental variable controlling the anoxygenic photosynthesis (van Gemerden and Mas, 1995). Therefore, a selective pressure for efficient light harvesting and maximum quantum yield exists in anoxygenic phototrophs. The same holds true for a few nichespecialized, deep-dwelling cyanobacteria. The ecological niches of green sulfur bacteria and Chromatiaceae show considerable overlap because both groups grow preferably or exclusively by photolithotrophic metabolism, using ambient sulfide as electron-donating substrate. Different species of the same group should be even more competitive. Besides differences in maintenance energy demand, in adaptation to low light intensities and metabolic flexibility, another important factor determining the species composition of phototrophic sulfur bacteria in their natural habitats is the spectral composition of underwater light. In the overlying layers, light is absorbed by water itself, dissolved yellow substance (gilvin), phytoplankton and inanimate particulates. The limited wavelength range available at great depth selects for species of anoxygenic phototrophic bacteria with complementary absorption spectra. In many lacustrine habitats, light absorption by phytoplankton exceeds that of gilvin or water itself (Kirk, 1983), and light of the blue green to green wavelength range reaches layers of phototrophic sulfur bacteria. Those Chromatiaceae which contain the carotenoid okenone (Fig. 7) dominated in 63% of the natural communities studied (van Gemerden and Mas, 1995). It was proposed that energy transfer from carotenoid antenna pigments to the reaction center is more efficient in okenone-forming strains than in other purple sulfur bacteria (Guerrero et al., 1986). In addition, the capability of gas vesicle formation, and the different kinetics of sulfide oxidation (see Coexistence of Phototrophic Sulfur Bacteria in this Chapter) appear to be of selective value for the colonization of pelagic habitats. Below accumulations of purple sulfur bacteria, the green-colored forms of the green sulfur bacteria dominate because of their superior capability to harvest the light reaching them, which has its spectrum shifted to a maximum intensity at 420–450 nm (Table 2) (Montesinos et al., 1997). In contrast, the brown-colored forms of the green sulfur bacteria dominate in lakes

CHAPTER 1.3

where the chemocline is located at depths greater than 9 m and in eutrophic lakes with a pronounced light absorption in the oxic zone. A similar niche separation occurs in the phototrophic consortia (see The Family Chlorobiaceae in Volume 7), which encompass greencolored or brown-colored epibionts (Overmann et al., 1999b). The ecological niche of the browncolored green sulfur bacteria may be attributed to their use of significantly lower light intensities than purple sulfur bacteria for phototrophic growth and to their lower maintenance energy requirements (see Light Absorption and Light Energy Transfer in Prokaryotes in this Chapter; The Family Chlorobiaceae in Volume 7). An extremely low-light adapted strain of the green sulfur bacterium Chlorobium phaeobacteroides has been isolated from the chemocline of the Black Sea located at an 80-m depth (Overmann et al., 1991a). This isolate (strain MN1) could grow at light intensities as low as 0.25 mmol quanta·m–2·s-1. In sedimentary environments with their particular optical properties (Fig. 6), the irradiance reaching anoxygenic phototrophic bacteria may be reduced to ¨1% of the surface value for light in the visible region, while >10% of the near infrared light is still available (Kühl and Jørgensen, 1992; see Light energy and the spectral distribution of radiation). As a consequence, the long wavelength Qy bands of bacteriochlorophylls are significant for light-harvesting in sediments, whereas light absorption of anoxygenic phototrophic bacteria in lakes is mediated by carotenoids and the Soret bands of bacteriochlorophylls. In microbial mats, the spectral quality of the scalar irradiance is strongly modified as it penetrates. The presence of populations of phototrophic microorganisms impose strong absorption signatures on the spectrum of the scalar irradiance (Jørgensen and Des Marais, 1988; Pierson et al., 1987). As a result of vertical niche separation, benthic microbial mats can consist of up to five distinctly colored layers that are formed (from the top) by diatoms and cyanobacteria, cyanobacteria alone, purple sulfur bacteria with bacteriochlorophyll a, purple sulfur bacteria with bacteriochlorophyll b, and green sulfur bacteria (Nicholson et al., 1987). In this vertical sequence different wavelength bands of red and infrared light (compare Table 2, Fig. 7) are successively absorbed by the different microbial layers (Pierson et al., 1990). Distinct blooms of bacteriochlorophyll b-containing anoxygenic phototrophic bacteria have been observed only in benthic habitats. Employing this pigment, the phototrophic Proteobacteria Blastochloris viridis, Blastochloris sulfoviridis, Thiocapsa pfennigii, Halorhodospira halochloris, Halorhodospira abdelmalekii harvest light of a wave-

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length range (1020–1035 nm), which cannot be exploited by any other photosynthetic organism. Until recently, no strain of anoxygenic photosynthetic bacteria was known that could absorb light in the wavelength range between 900 and 1020 nm. Because of the prevalence of infrared radiation in the anoxic layers of microbial mats and the strong competition for this wavelength region, bacteria containing other types of photosynthetic antenna complexes would have a high selective advantage. Recently, the aProteobacterium Rhodospira trueperi was isolated, which contains bacteriochlorophyll b in a light-harvesting complex with a maximum absorption at 986 nm (Pfennig et al., 1997). Employing a selective enrichment strategy, the a-Proteobacterium Roseospirillum parvum could be isolated which harbors another new type of photosynthetic antenna complex. Here, bacteriochlorophyll a is the light-harvesting pigment and in vivo exhibits an absorption maximum at 911 nm (Glaeser and Overmann, 1999, Fig. 7). Both isolates originate from benthic microbial mats, indicating that the diversity of pigment-protein complexes in Proteobacteria is higher than previously assumed. The variation in the in vivo absorption spectra of the same pigment must be the result of differences in binding to light-harvesting proteins. In contrast, changes in the absorption spectra of the light-harvesting complex of green sulfur bacteria are the result of chemical alterations (e.g., methylation) of the pigment molecules (Bobe et al., 1990) because pigment-pigment interactions dominate in the chlorosomes (see Light Absorption and Light Energy Transfer in Prokaryotes in this Chapter). Because methanogenesis is the predominant pathway of terminal degradation in rice fields, Heliobacteriaceae probably compete with the photoheterotrophic purple nonsulfur bacteria in their natural environment (Madigan and Ormerod, 1995). Owing to the presence of bacteriochlorophyll g, Heliobacteriaceae take advantage of a wavelength region of the electromagnetic spectrum, which is not absorbed by other phototrophic bacteria. As a result of the small and fixed size of the photosynthetic antenna (see Light Absorption and Light Energy Transfer in Prokaryotes in this Chapter), these bacteria are adapted to higher light intensities than other anoxygenic phototrophic bacteria (ª1,000 mol quanta·m–2·s-1). In addition to the capacity of absorbing light in the long wavelength range, metabolic flexibility is of highly selective value for the colonization of benthic habitats with their high fluctuations in oxygen and sulfide concentrations (see Chemotrophic growth with O2). However, the composition of communities of phototrophic sulfur bacteria is not solely deter-

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mined by competition. The simultanous presence of green sulfur bacteria and Chromatiaceae possibly is also based on syntrophic interactions (see Coexistence of Phototrophic Sulfur Bacteria in this Chapter). Coexistence of Phototrophic Bacteria Within the Chromatiacea the small-celled genus Chromatium species exhibit a considerably greater metabolic flexibility than the large-celled species (see Carbon Metabolism and Chemotrophic Growth with O2 in this Chapter). In addition, small-celled species like Allochromatium vinosum have a higher growth affinity for sulfide. Based on these pure culture data, it is therefore unexpected that large-celled forms in fact dominate in natural ecosystems. The largecelled Chromatium weissei oxidizes sulfide twice as fast as the small-celled Allochromatium vinosum. Whereas the former preferentially oxidizes sulfide to zero-valent sulfur, the latter oxidizes a larger fraction directly to sulfate. Under fluctuating conditions as they occur in the chemocline of lakes, Chromatium weissei is capable of rapidly oxidizing sulfide at the onset of illumination, thereby accumulating zero-valent sulfur. During the remaining light period and because of its higher affinity for sulfide, Allochromatium vinosum utilizes most of the sulfide. Continuous cocultures of both species have thus been established by illumination in light-dark cycles (van Gemerden, 1974). Furthermore, stable coexistence of two organisms is feasible in the presence of two substrates for which the two competitors have complementary affinities. Stable syntrophic interactions can be established in laboratory cocultures of purple sulfur (Allochromatium vinosum) and green sulfur bacteria (Chlorobium limicola f.sp. thiosulfatophilum; van Gemerden and Mas, 1995). Because of its higher affinity, the green sulfur bacterium oxidizes sulfide to zero-valent sulfur. The extracellular sulfur is mobilized as polysulfide, which can be used instantaneously as electron donor of the purple sulfur bacterium. The presence of sulfide inhibits the green sulfur bacterium from using polysulfide (see The Family Chlorobiaceae in Volume 7). Sulfide and polysulfide thus are the mutual substrates for the two different phototrophic sulfur bacteria. Purple and green sulfur bacteria also have complementary affinities for sulfide and acetate (see Carbon metabolism). Accordingly, stable continuous cocultures of Chlorobium phaeobacteroides and Thiocapsa roseopersicina can be established (Veldhuis and van Gemerden, 1986). Interactions Between Phototrophic Sulfur Bacteria and Chemotrophic Bacteria A considerable number of strains of Chromatiacae is

CHAPTER 1.3

capable of switching to a chemolithotrophic growth mode after prolonged incubation in the presence of molecular oxygen (see Chemotrophic Growth with O2 in this Chapter). Under these conditions, purple sulfur bacteria compete with colorless sulfur bacteria like Thiobacillus spp. Compared to thiobacilli, the purple sulfur bacterium Thiocapsa roseopersicina attains a higher growth yield under chemolithotrophic conditions (De Wit and van Gemerden, 1987a). However, the growth affinity for sulfide of the colorless sulfur bacteria is up to 47 times higher than that of Chromatiacae (De Wit and van Gemerden, 1987b; van Gemerden and Mas, 1995). Therefore Chromatiacae growing exclusively by chemolithotrophy would be rapidly outcompeted by colorless sulfur bacteria. Culture experiments indicate that Thiocapsa roseopersicina, a typical inhabitant of laminated microbial mats in temperate environments, can replenish its photosynthetic pigments during anoxic periods in the dark, thereby maintaining a phototrophic growth mode also during the subsequent oxic light period (De Wit and van Gemerden, 1990b). Based on microelectrode measurements, purple sulfur bacteria in marine microbial mats of the North Sea barrier islands are exposed to oxygen during most of the day, whereas anoxic conditions prevail during the night (De Wit et al., 1989). Thus, the anoxygenic phototrophs cannot grow during the night and face competition for sulfide by colorless sulfur bacteria during the day. Because of their higher affinity for sulfide, the latter would be expected to outcompete phototrophically growing purple sulfur bacteria. In cocultures of Thiocapsa roseopersicina and Thiobacillus thioparus, sulfide is indeed entirely used by the colorless sulfur bacterium in the presence of oxygen. If oxygen concentrations are limiting, however, sulfide is oxidized incompletely by the chemolithotroph and soluble zero-valent sulfur formed (either as polysulfide or polythionates) that in turn is used by the purple sulfur bacterium for phototrophic growth (van den Ende et al., 1996). Both diurnal fluctuations between oxic light and anoxic dark periods and syntrophism based on sulfur compounds may permit a stable coexistence of these groups and explain their simultaneous presence in natural microbial mats. Stable associations can be established between green sulfur bacteria and sulfur- or sulfatereducing bacteria (see The Family Chlorobiaceae in Volume 7; Interactions with Chemotrophic Bacteria in this Chapter). These associations are based on a cycling of sulfur compounds but not carbon (see Significance of Anoxygenic Photosynthesis for the Pelagic Carbon and Sulfur Cycles in this Chapter). The simultaneous growth of both types of bacteria is fueled by the

CHAPTER 1.3

oxidation of organic carbon substrates and light. In a similar manner, cocultures of Chromatiaceae with sulfate-reducing bacteria have been established in the laboratory (van Gemerden, 1967). The most spectacular type of association involving phototrophic bacteria is represented by the phototrophic consortia. These consortia consist of green sulfur bacterial epibionts that are arranged in a regular fashion around a central chemotrophic bacterium. A rapid signal transfer exists between the two partners and permits phototrophic consortia to scotophobotactically accumulate at preferred light intensities and wavelengths. In this association, the immotile green sulfur bacteria attain motility like purple sulfur bacteria. The high numbers of phototrophic consortia found in many lakes indicate that this strategy must be of high competitive value under certain environmental conditions. A commensal relationship may exist between coccoid epibiotic bacteria and the purple sulfur bacterium Chromatium weissei (Clarke et al., 1993). This unidentified epibionts attaches to healthy cells but does not form lytic plaques on lawns of host cells like the morphologically similar parasite Vampirococcus (see Significance of Bacteriophages and Parasitic Bacteria in this Chapter). Possibly, the epibiont grows chemotrophically on carbon compounds excreted by the purple sulfur bacterium. A syntrophic interaction between cyanobacteria and sulfate-reducing bacteria appears to exist in microbial mats where both types of microorganisms occur in close spatial proximity, if not intermixed with each other. In these ecosystems, the excretion of organic carbon substrates by cyanobacteria may provide the electrondonating substrates for sulfate-reducing bacteria (Jørgensen and Cohen, 1977; Skyring and Bauld, 1990; Fründ and Cohen, 1992). The glycolate produced by photorespiration (Fründ and Cohen, 1992), as well as the formate, acetate and ethanol produced by glycogen fermentation (Moezelaar and Stal, 1994) most likely are the substrates excreted by cyanobacteria. Despite a pronounced limitation of sulfate reduction by carbon substrates (Overmann et al., 1996; Overmann, 1997), no close syntrophic relationship was found between purple sulfur and sulfate-reducing bacteria in a meromictic lake. In this specific environment degradation of biomass by the entire anaerobic food chain rather than excretion of small carbon molecules and their direct utilization by sulfate-reducing bacteria provides the electron-donating substrates for sulfate-reducing and sulfur-reducing bacteria. Symbioses Between Phototrophic Bacteria and Eukaryotes Only one example is known for an intracellular symbiosis of anoxygenic pho-

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totrophic bacteria with an eukaryotic organism. The ciliate Strombidium purpureum inhabits the photic zone of sulfide-containing marine sands and harbors 200–700 purple endosymbionts. Symbionts are arranged along the periphery of the host cell and contain intracellular tubular or vesicular membranes, bacteriochlorophyll a and spirilloxanthin (Fenchel and Bernard, 1993a; Fenchel and Bernard, 1993b). The ciliate shows a photosensory behavior, accumulating at wavelength that corresponds to the absorption maxima of the endosymbionts. It has been suggested that the intracytoplasmic purple bacteria increase the efficiency of the fermentative host by using its end products for anoxygenic photosynthesis. Furthermore, respiration of the bacteria may protect the host against oxygen toxicity. In the course of evolution, Cyanobacteria have entered into symbiotic associations with a multitude of organisms (Schenk, 1992). Besides all eukaryotic phototrophs, from microalgae to Sequoia sempervirens, which have intracellular cyanobacterial symbioses, the most common extracellular symbioses of nonheterocystous cyanobacteria are in the form of cyanolichens and involve the unicellular genera Chroococcidiopsis, Gloeocapsa, “Chroococcus,” and Gloeothece, as well as members of the genera Nostoc, Calothrix, Scytonema, Stigonema, and Fischerella as photobionts. Heterocystous cyanobacteria in the genus Nostoc form extracellular symbioses with liverworts and hig her plants (Cycads, duckweed). Anabaena enters in symbiosis with water ferns of the genus Azolla. Prochloron strains, large-celled Synechocystis and small-celled Acaryochloris marina are known from extracellular symbioses with ascidians in tropical or subtropical marine waters; Prochloron is found as ectosymbiont on the marine didemnid ascidian Lissoclinum patella (Lewin and Withers, 1975). Extracellular symbioses of the Pseudanabaena-like “Konvophoron” occur in Mediterranean invertebrates. Finally, intracellular symbioses of nonheterocystous cyanobacteria are known with tropical sponges (“Aphanocapsa”, Oscillatoria, Synechocystis, Proc hloron), with green algae (Phormidium) and dinoflagellates (unidentified). Heterocystous cyanobacteria occur intracellularly in oceanic diatoms of the genera Hemiaulus and Rhizosolenia (and the cyanobacterium Richelia intracellularis). The cyanobacterial symbiont consists of a short cell filament with a terminal heterocyst (Mague et al., 1977). The numbers of filaments varies with host species. Nostoc thrives intracellularly in Trifolium (clover) and also in the terrestrial non-lichenic fungus Geosiphon pyriforme. With the notable exception of lichenic photobionts, many symbiotic cyanobacteria have resisted cultivation in spite of continued efforts.

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Significance of Bacteriophages and Parasitic Bacteria In addition to grazing, light and nutrient limitation, cyanophage infection of cyanobacteria may be a significant factor limiting primary productivity in the marine environment. However, because of inactivation by solar radiation and resistence of the host cells, the role of cyanophages has remained unclear (Bergh et al., 1989; Proctor and Fuhrman, 1990; Suttle et al., 1990; Suttle et al., 1993; Waterbury and Valois, 1993). Several bacteria have been discovered that attack phototrophic bacteria (Guerrero et al., 1986; Nogales et al., 1997). Vampirococcus attaches to the cell surface of Chromatium spp. where it divides, forming chains of up to three cells. Concomitantly, the cytoplasm of the host cell appears to be degraded. Daptobacter penetrates the cell envelope and divides intracellularly by binary fission. In contrast to Vampirococcus, Daptobacter has been cultivated in the absence of the host and grows by fermentative metabolism. Bdellovibrio has a broad host range, and under laboratory conditions attacks also purple sulfur bacteria. Bdellovibrio forms daughter cells by multiple division in the periplasmic space of the host cell. The Gramnegative chemotrophic bacterium Stenotrophomonas maltophilia is a non-obligatory parasite of green sulfur bacteria, which causes cell lysis and ghost formation (Nogales et al., 1997). Its host range is not limited to green sulfur bacteria. The presence of parasitic bacteria in water samples becomes evident by the formation of lytic plaques on lawns of host bacteria (Esteve et al., 1992; Nogales et al., 1997). Up to 94% of the cells of phototrophic sulfur bacteria may be infected by parasitic bacteria in natural samples. Since infection is largely limited to nongrowing cells, the impact of parasitism on populations of phototrophic sulfur bacteria appears to be limited (van Gemerden and Mas, 1995).

Evolutionary Considerations Porphyrins are found in all organisms from archaebacteria through plants to animals, and are indispensable as prosthetic groups for energy conservation. In contrast, the partially reduced derivates of porphyrins, the (bacterio)chlorophylls, are synthesized by members of only a few bacterial divisions (Fig. 1). This indicates that the capability for synthesis of porphyrins is a very ancient trait, whereas only a few prokaryotes acquired the capability to form photosynthetic pigments. Photosynthesis requires the presence of various complex protein structures and cofactors, and thus the expression of a large number of different genes (see Photosynthetic Gene Cluster in this Chapter). Previously, it had there-

CHAPTER 1.3

fore appeared justified to consider all phototrophic prokaryotes as a monophyletic group only distantly related to nonphototrophic bacteria (Pfennig and Trüper, 1974; Trüper and Pfennig, 1978). Two lines of evidence have been used to recon struct the evolution of photosynthesis. Fossil Evidence The oldest fossils of microorganisms have been dated back to the early Archaean (3.8 billion years ago) and may represent remains of cyanobacteria (Awramik, 1992). They consist of chemical fossils and stromatolites that have been detected especially in sedimentary rocks of the Pilbara region, Western Australia, and the Barberton Mountain Land, South Africa. Stromatolites are laminated convex domes and columns of cm to dm size and have been found in 3.5 to 0.8 billion year old rocks. Although scarce in biosynthetic molecular skeletons, the insoluble, high-molecular-weight organic matter (kerogen) contains isotopic evidence for autotrophic carbon fixation. The ratio of stable carbon isotopes (d13C values) are in the range of –35.4 to –30.8∞/oo, which is typical for CO2-carbon fixed by the ribulose-1,5bisphosphate cycle (Hayes et al., 1983). In addition, the se ancient sediments contain laminated domes and columns of cm to dm size, which in analogy to extant stromatolites have been interpreted as organosedimentary structures produced by the trapping, binding, and precipitation activity of filamentous microorganisms, most likely cyanobacteria. Alternatively, it has been proposed that anoxygenic photosynthetic bacteria and not the oxygenic cyanobacteria formed the oldest stromatolites. Based on the phylogenetic analysis of the 16S rRNA gene sequence (Oyaizu et al., 1987) and the ecophysiology (Ward et al., 1989) of the filamentous green photosynthetic bacterium Chloroflexus aurantiacus, similar anoxygenic phototrophic bacteria may be the more likely candidate microorganisms that built the most ancient stromatolites. However, according to analyses of the nucleotide sequences of its reaction center polypeptides and primary sigma factor (see Molecular Evidence in this Chapter), Chloroflexus aurantiacus does not represent a deep branch of bacterial evolution. Gypsum layers within the supposed stromatolites have been interpreted as indicators of sulfide oxidation by either anoxygenic phototrophs or colorless sulfur-oxidizing bacteria (Awramik, 1992). However, similar structures have been discovered in lacustrine, and thus sulfur-depleted, settings with little input of allochthonous organic carbon (Buick, 1992). Therefore, at least some 2.7 billion year-old stromatolites are more likely to have harbored oxygenic cyanobacteria. Taken together with the fossil evidence, this would indi-

CHAPTER 1.3

cate that diversification of the major groups of phototrophic microorganisms did occur during the early Archaean (Awramik, 1992). Because of the indefinite character of the fossil evidence, 16S rRNA sequences and components of the photosynthetic apparatus of the different photosynthetic prokaryotes have been used to gain additional insight into the evolution of photosynthesis. Molecular Evidence Chlorophyll-based photosystems are only found in the Bacteria and chloroplasts, suggesting that this type of energy conversion originated in the bacterial lineage after the divergence of Archaea and Eukarya. So far, photosynthetic species have not been discovered in the very early lineages of the bacterial radiation (e.g., the thermophilic oxygen reducers and Thermotogales; Fig. 1). Because most species of these lineages are chemolithotrophic, it has been proposed that chemolithoautotrophy preceded phototrophy during the evolution of the Bacteria (Pace, 1997). This conclusion is supported by the fact that in phylogenetic trees based on protein sequences of elongation factor EF-Tu and the b-subunit of ATP synthase, only the Aquificales and Thermotogales branch deeper than the majority of the bacterial divisions, while the Chloroflexus subdivision does not (Stackebrandt et al., 1996), thus indicating that Chloroflexus does not represent the descendant of a more ancient ancestor than other phototrophic bacteria. At present, five of the known bacterial lineages comprise phototrophic species (Fig. 1, see Taxonomy of Phototrophy among Prokaryotes in this Chapter). Based on 16S rRNA sequences, extant phototrophic species of different lineages are only very distantly related to each other. Furthermore, one lineage, the Chloroflexus subgroup, represents an early branch in the evolution of the Bacteria. Given the complexity of the photosynthetic apparatus, it is unlikely that photosynthesis has evolved more than once during the evolution of the domain Bacteria (Woese, 1987). The phylogenetic analysis indicates that either an early ancestor of most known bacteria had acquired the capacity for photosynthetic growth (Stackebrandt et al., 1988) or, alternatively, that the genes coding the photosynthetic apparatus were transferred laterally between phylogenetically distant bacteria. The evidence for the various scenarios of the evolution of bacterial photosynthesis is discussed in the present section. Originally, it had been proposed (Oparin, 1938; Gest and Schopf, 1983a) that anaerobic, heterotrophic prokaryotes capable of fermenting hexose sugars were among the earliest life forms and that electron transport and photosynthesis

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evolved as a response to the depletion of organic nutrients from the primordial soup. Based on one hypothesis (the Granick hypothesis; Granick, 1965), the biosynthetic pathway of photosynthesis pigment molecules may be taken as a recapitulation of evolution such that compounds with shorter biosynthetic pathways reflect the more ancestral state. The synthesis of bacteriochlorophyll requires one additional enzymatic reduction than that of chlorophyll. Because chlorophyll precedes bacteriochlorophyll in the biosynthetic pathway, the former should have existed earlier in nature. It has been proposed (Pierson and Olson, 1989) that a non-oxygenic photosynthetic ancestor containing chlorophyll a and the two types of reaction centers evolved prior to the major radiation event of the Bacteria. During the subsequent radiation, oxygen evolution appeared in one line of descent whereas either the quinone or the FeS-type photosystem was lost in other lineages, concomitant with the emergence of the different bacteriochlorophylls. Besides avoiding an a priori lateral gene transfer of the complete photosynthetic gene cluster, this Pierson-Olson hypothesis takes into account the ecological conditions of the early biosphere in which the absence of oxygen and ozone caused a predominance of radiation in the blue and UV wavelength range, which in turn would render the red-shifted absorption maxima of bacteriochlorophylls of little selective advantage (Boxer, 1992). As an argument against the Granick and Pierson-Olson hypotheses, several types of phototrophic bacteria that would be expected are apparently missing in nature. As an example, anoxygenic chlorophyll-containing forms have never been found, although it has been argued that the 8-hydroxychlorophyll-containing Heliobacteriaceae represents this type inasmuch as bacteriochlorophyll g is easily converted to chlorophyll a by oxidation. Bacteriochlorophylls occur in both types of reaction centers, the pheophytin-type (Proteobacteria, Chloroflexus) and the FeS-type. This could indicate that the presence of bacteriochlorophyll represents a primitive trait. The chlorophyll-first hypothesis postulates that bacteriochlorophyll has replaced chlorophyll independently in at least three different bacterial lineages. Chlorophyll, however, is presently only found in oxygen-evolving organisms of the phylum Cyanobacteria whic h, based on 16S rRNA sequence comparison, represents the most recently evolved group of phototrophic bacteria (Woese, 1987, Fig. 1). Cyanobacteria contain two different photosystems and thus have the most complex photosynthetic apparatus. In addition, the much higher complexity of the oxygen-evolving PSII of oxygenic phototrophic organisms may imply that it

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appeared later than the other photosystems during evolution. As another argument against the PiersonOlson hypothesis, chlorophyll itself should have been of little selective advantage in Earth’s early biosphere and it has been proposed that quinone-iron complexes represented the first photosynthetic unit (Boxer, 1992). In contrast to the complex porphyrin pigments, quinones can form spontaneously from acetyl thioesters (Hartmann, 1992). Furthermore, the discrepancy between the presence of chlorophyll exclusively in the most highly evolved bacteria and its shorter biosynthetic pathway may be explained by the finding that the chlorin reductase, which catalyzes the additional step of the biosynthetic pathway for bacteriochlorophyll, is phylogenetically older than the enzyme (protochlorophyllide reductase) that catalyzes the preceeding step. This enzyme is present in both the chlorophyll- and bacteriochlorophyll-containing bacteria (Burke et al., 1993). An ancien t reductase may have been able to perform both, the reduction of protochlorophyllide and of chlorin, such that bacteriochlorophyll was the photochemically active pigment in the last common ancestor of all extant phototrophic bacteria. An analysis of the distribution of the different types of reaction centers among the different bacterial phyla and the amino acid sequences of reaction center proteins (Blankenship, 1992) provides an alternative hypothesis for the evolution of photosynthesis, namely the possibility of lateral transfer of photosynthesis genes. Both the pheophytin/quinone and the FeS-type reaction centers are found in phylogenetically distant groups (e.g., a pheophytin/quinone reaction center in Chloroflexus and phototrophic members of the a-Proteobacteria). Even more significantly, a phylogenetic analysis of the amino acid sequences of pheophytin-type reaction center polypeptides from the three different bacterial lineages Chloroflexaceae, cyanobacteria and aProteobacteria indicated that the reaction center of Chloroflexus aurantiacus is more closely related to that of phototrophic members of the a-Proteobacteria than to the PSII reaction center of cyanobacteria (Blankenship, 1992). Thus the reaction center of Chloroflexus must have evolved after (and not prior to) the divergence of the D1/D2 branch from the L/M line of descent. Another essential component of the photosynthetic apparatus of Chloroflexus and green sulfur bacteria are the light-harvesting chlorosomes. Based on amino acid sequence comparison of protein constituents, chlorosomes of both groups have a common evolutionary origin (Wagner-Huber et al., 1988). Similarly, a comparison of the amino acid sequences of the group 1 s70 primary sigma factor also has dem-

CHAPTER 1.3

onstrated a close relationship to the green sulfur bacteria with respect to this component of the central housekeeping function (Gruber and Bryant, 1998). Other features of Chloroflexus aurantiacus appear to be unique (like the lipid and carotenoid composition), or ancient (like the hydroxypropionate pathway of CO2-fixation). Recently, the activity of the key enzymes of this pathway have been reported for some archaea (Menendez et al., 1999) such that Chloroflexus aurantiacus seems to represent a “chimeric” organism. Based on the most parsimonious assumption that homodimeric reaction centers are ancestral to homodimeric ones, the reaction centers of green sulfur bacteria and Heliobacteriaceae would resemble most the reaction center of the ancestor of all extant bacteria. It has been hypothesized (Gruber and Bryant, 1998) that the reaction center of Chloroflexus aurantiacus was acquired by a recent lateral gene transfer event that may have replaced a type I reaction center with a type II (FeS) reaction center, whereas other features like primary sigma factor or chlorosomes still reflect the common descent of Chloroflexus and the green sulfur bacteria. Alternatively, it has been suggested that transfer of the genetic information of the relatively simple chlorosomes occurred after the evolution of the two classes of reaction centers and that the green sulfur bacteria represent a relatively modern evolutionary invention (Stackebrandt et al., 1996). The presence of two homologous polypeptides in all known reaction centers would suggest a single gene duplication event in an early ancestor of all phototrophic bacteria. As an additional result of the phylogenetic analysis of the amino acid sequences of pheophytin-type reaction center polypeptides from the three different bacterial lineages (Chloroflexaceae, cyanobacteria and a-Proteobacteria; Blankenship, 1992), the most likely occurrence of two independent gene duplications is suggested—one leading to the reaction center of PSII in cyanobacteria and green plants (polypeptides D1 and D2) and another to the reaction center of Chloroflexus and purple nonsulfur bacteria (polypeptides L and M). Another, third, independent gene duplication has to be assumed during the evolution of the FeS-type reaction center. The reason for the paraphyletic development of the three lineages may be a functional advan tage of dimeric reaction centers over monomeric ones. Yet another evolutionary scenario for photosynthetic reaction centers (Vermaas, 1994) has been based on the finding that the sixth membrane-spanning region of the heliobacterial (FeS- or PSI-type) reaction center shows a great similarity to the sixth membrane-spanning

CHAPTER 1.3

region of the CP47 antenna polypeptide of (the quinone-type) PSII, and the preceeding Nterminal five hydrophobic regions still show significantly greater similarity to CP47 (and to another PSII antenna protein, CP43) than to the respective portion of PSI. According to this model, an ancestral homodimeric antenna/reaction center complex comprised 11 putative transmembrane regions and contained two quinones and an Fx-type Fe4S4 iron-sulfur center. Relatively few modifications may have led to the homodimeric complex of green sulfur bacteria and Heliobacteriaceae, whereas a gene duplication event and divergent evolution led to the heterodimeric PSI. As a para llel line of descent, splitting of the ancestral reaction center complex into a reaction center and a separate antenna protein may have occurred. Operon duplication, loss of the FeS, and divergent evolution are assumed to have resulted in two separate lineages. By association with an additional watersplitting enzyme system, PSII was formed. In contrast, the separate antenna polypeptide was lost and replaced by a modified antenna complex (light-harvesting I) during evolution of the reaction center of Proteobacteria and Chloroflexus. Significantly, however, this theory does not explain the occurrence of the quinone-type reaction center in these latter two groups, which are phylogenetically very distant. In addition, the combination of a reaction center typical for Proteobacteria with an antenna structure characteristic for green sulfur bacteria would still need to be explained by lateral gene transfer of either of the two components. Based on the obvious discrepancy between the phylogeny of ribosomal RNA and reaction center proteins, the hypothesis of lateral transfer of photosynthesis genes between distantly related groups of bacteria has been put forward. Lateral gene transfer as yet seems to provide the simplest explanation for the distribution pattern of photosynthesis genes within the bacterial radiation (Blankenship, 1992; Nagashima et al., 1993; Nagashima et al., 1997). Such a lateral gene transfer would encompass reaction center structural genes, genes coding for other electron transfer proteins, and genes needed for the biosynthesis of pigments and cofactors. In purple nonsulfur bacteria the majority of these genes indeed form a single cluster of 46 kb (which does not encompass the genes for the light-harvesting II complex, however; Bauer and Bird, 1996; Wellington et al., 1992; Yildiz et al., 1992). The genetic organization may be taken as evidence for lateral gene transfer as the cluster represents only ~1.3% of the total genome size. It should be mentioned, however, that clustering of most photosynthesis genes may also be due to structural or regulatory constraints. Supporting the

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latter argument (Yildiz et al., 1992), photosynthesis genes in a-Proteobacteria are transcriptionally coupled in superoperons involving overlapping transcripts. The particular genetic organization is the prerequisite for adaptation of the cells to changing light intensity (see Genetic Regulation in Response to Light in this Chapter) and oxygen tension (see Genetic Regulation by O2 in this Chapter). Therefore a selective pressure may exist to retain the linkage order and would make the genetic organization of the photosynthesis genes less suitable for phylogenetic inference. Furthe rmore, the high correlation between the phylogenetic trees for 16S rRNA and cytochrome c in phototrophic members of the a-Proteobacteria has been taken as evidence that a lateral transfer of photosynthesis genes did not occur at least within this phylogenetic group (Woese et al., 1980). Thus, the presence of reaction centers in aerobic bacteriochlorophyllcontaining a-Proteobacteria may represent an atavistic trait, and the genes coding the reaction center might have been lost frequently during the evolution of aerobic representatives in this group (Stackebrandt et al., 1996). Because the pigment composition of the oxygenic photosynthetic “Prochlorophytes” is very similar to that of green plant chloroplasts, and like the latter “Prochlorophytes” have appressed thylakoid membranes, it has been proposed that the chloroplasts of green plants evolved from an endosymbiotic “prochlorophyte” (van Valen and Maiorana, 1980; Lewin, 1981). In contrast to the other oxygenic phototrophs, Prochlorococcus contains divinyl isomers of chlorophylls a and b, and a- instead of b-carotene (Chrisholm et al., 1992; Goericke and Repeta, 1992). However, based on sequence comparison of 16S rRNA (Urbach et al., 1992) and the rpoC1 (Palenik and Haselkorn, 1992) genes, the three known prochlorophyte lineages (Prochloron, Prochlorothrix, and Prochlorococcus) are no direct ancestors of chloroplasts. In addition, these analyses revealed that “Prochlorophytes” most likely are of polyphyletic origin and that the use of chlorophyll b as additional light-harvesting pigments must have developed at least four times during evolution. In this case, too, a horizontal transfer of the respective biosynthesis genes could be invoked to explain the distribution pattern of chlorophyll b among the different members of the cyanobacterial division (Palenik and Haselkorn, 1992). Immunological studies and differences in the chlorophyll a/chlorophyll b ratio of the antennae isolated from different “Prochlorophytes” indicate that the capacity to bind chlorophyll b arose several times and independently from the cyanobacterial ancestors, and thus confirm the results of s equence comparisons of the 16S rRNA and rpoC1 genes.

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Strauß, G., Fuchs, G. 1993. Enzymes of a novel autotrophic CO2 fixation pathway in the phototrophic bacterium Chloroflexus aurantiacus, the hydroxypropionate cycle. Eur J Biochem 215:633–643. Suttle, C. A., Chan, A. M., Cottrell, M. T. 1990. Infection of phytoplankton by viruses and reduction of primary productivity. Nature 347:467–469. Suttle, C. A., Chan, A. M., Feng, C., Garza, D. R. 1993. Cyanophages and sunlight: A paradox. In: Guerrero R, Pedrós-Alió C (Eds.) Trends in Microbial Ecology. Spanish Society for Microbiology. Barcelona, 303–307. Suwanto, A., Kaplan, S. 1989. Physical and genetic mapping of the Rhodobacter sphaeroides 2.4.1 genome: genome size, fragment identification, and gene localization. J Bacteriol 171:5840–5849. Suzuki, J. Y., Bauer, C. E. 1995. A prokaryotic origin for lightdependent chlorophyll biosynthesis of plants. Proc Natl Acad Sci USA 92:3749–3753. Swoager, W. C., Lindstrom, E. S. 1971. Isolation and counting of Athiorhodaceae with membrane filters. Appl Microbiol 22:683–687. Takahashi, K., Wada, E., Sakamoto, M. 1990. Carbon isotope discrimination by phytoplankton and photosynthetic bacteria in monomictic Lake Fukami-ike. Arch Hydrobiol 120:197–210. Tanada, T., Kitadokoro, K., Higuchi, Y., INaka, K., Yasui, A., Deruiter, P. E., Eker, A. P. M., Miki, K. 1997. Crystal structure of DNA photolyase from Anacystis nidulans. Nature Struct. Biol 4:887–891. Thauer, R. K., Jungermann, K., Decker, K. 1977. Energy conservation in chemotrophic anaerobic bacteria. Bacteriolog Rev 41:100–180. Theroux, S. J., Redlinger, T. E., Fuller, R. C., Robinson, S. J. 1990. Gene encoding the 5.7-kilodalton chlorosome protein of Chloroflexus aurantiacus: regulated message levels and a predicted carboxy-terminal protein extension. J Bacteriol 172:4497–4504. Thomas, R. H., Walsby, A. E. 1985. Buoyancy regulation in a strain of Microcystis. J Gen Microbiol 131:799– 809. Thorne, S. W., Newcomb, E. H., Osmond, C. B. 1977. Identification of chlorophyll b in extracts of prokaryotic algae by fluorescence spectroscopy. Proc Natl Acad Sci 74:575–578. Trüper, H. G., Pfennig, N. 1978. Taxonomy of the Rhodospirillales. In: Clayton RK, Sistrom WR (Eds.) The photosynthetic bacteria. Plenum Publishing Corp.. New York, 19–27. Trüper, H. G., Pfennig, N. 1981. Characterization and identification of the anoxygenic phototrophic bacteria. In: Starr MP, Stolp H, Trüper HG, Balows A, Schlegel HG (Eds.) The prokaryotes: A handbook on habitats, isolation and identification of bacteria. Springer. New York, 299–312. Turner, S. 1887. Molecular systematics of oxygenic photosynthetic bacteria. Plant Syst Evol 11:13–52. Urakami, T., Komagata, K. 1984. Protomonas, a new genus of facultatively methylotrophic bacteria. Int J Syst Bacteriol 34:188–201. Urbach, E., Robertson, D. L., Chrisholm, S. W. 1992. Multiple evolutionary origins of prochlorophytes within the cyanobacterial radiation. Nature 355:267–270. Utkilen, H. C., Skulberg, O. M., Walsby, A. E. 1985. Buoyancy regulation and chromatic adaptation in planktonic Oscillatoria species: alternative strategies for optimizing

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CHAPTER 1.3 waters of the Juan de Fuca Ridge in the Pacific Ocean. Appl Environ Microbiol 64:337–341. Zehnder, A. J. B., Stumm, W. 1988. Geochemistry and biogeochemistry of anaerobic habitats. In: Zehnder AJB (Eds.) Biology of Anaerobic Microorganisms. WileyLiss. New York, 1–38. Zevenboom, W., Mur, L. R. 1984. Growth and photosynthetic response of the cyanobacterium Microcystis aeruginosa in relation to photoperiodicity and irradiance. Arch Microbiol 139:232–239.

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Zuber, H., Cogdell, R. J. 1995. Structure and organization of purple bacterial antenna complexes. In: Blankenship RE, Madigan MT, Bauer CE (Eds.) Anoxygenic Photosynthetic Bacteria. Kluwer Academic Publishers. Dordrecht, The Netherlands. 315–348. Zucconi, A. P., Beatty, J. T. 1988. Posttranscriptional regulation by light of the steady state levels of mature B800850 light-harvesting complex in Rhodobacter capsulatus. J Bacteriol 170:877–882.

Prokaryotes (2006) 2:86–101 DOI: 10.1007/0-387-30742-7_4

CHAPTER 1.4 ehT

c i boreanA

yW a

fo

The Anaerobic Way of Life RUTH A. SCHMITZ, ROLF DANIEL, UWE DEPPENMEIER AND GERHARD GOTTSCHALK

Introduction Molecular oxygen in appreciable amounts is found only in those areas on earth that are in direct contact with air or are inhabited by organisms carrying out oxygenic photosynthesis. The solubility of oxygen in water is low. In equilibrium with air at 1.013 bar and at 20∞C, pure water will contain approximately 9 mg/liter of dissolved oxygen. In aqueous systems, aerobic organisms rapidly consume dissolved oxygen, so that deeper layers of many waters and soils (especially if they are rich in organic compounds), as well as mud and sludge, are practically anaerobic. Nevertheless, these areas are inhabited by numerous organisms that fulfill the important ecological role of converting insoluble organic material to soluble compounds and gases that can circulate back into aerobic regions. Other important anaerobic habitats are the rumen, the intestinal tract, and man-made anaerobic digestors of sewage treatment plants. Anaerobic prokaryotes that can live in the above-mentioned environments are either phototrophs, which, of course, can only flourish if light is available, or chemotrophs. With respect to their relationship to aerobic metabolism, three groups of organisms capable of growth in an anaerobic environment can be identified: 1. Organisms that are aerobes but can use alternate electron acceptors such as nitrate or nitrite when exposed to an anaerobic environment. The electron transport from NADH to these acceptors is coupled to the phosphorylation of ADP, as is the electron transport to oxygen. 2. Organisms that are facultative aerobes. The enterobacteria are the most prominent representatives of this group. These organisms grow as typical aerobes in the presence of oxygen; in its absence, they carry out fermentations. 3. Obligately anaerobic bacteria that are characterized by the inability to synthesize a respiratory chain with oxygen as terminal electron acceptor. They are restricted to life without oxygen.

The diversity of microorganisms able to thrive under anaerobic conditions is overwhelming. Up to now more than 200 genera of obligate anaerobic microorganisms have been described. Obligate anaerobes are found in all three domains. The eukaryotes are represented by anaerobic fungi, ciliates and flagellates, the archaea by the methanogens, which comprise 23 genera, and by the most hyperthermophilic genera Pyrolobus, Pyrodictium and Pyrococcus. Most genera of the obligate anaerobes belong to the bacteria. Especially prominent are the 32 genera characterized by their ability of dissimilatory reduction of sulfate, sulfite or sulfur. Spore formers are well represented, e.g., by the genera Clostridium, Sporomusa, Desulfotomaculum, Moorella and Thermoanaerobacterium. There are halophiles such as the genera Haloanaerobacter and Sporohalobacter and alkaliphiles like Anaerobranca. A few genera comprise more than a dozen species: Bacteroides, Bifidobacterium and Clostridium (the genus which by far contains the most species), Desulfotomaculum, Desulfovibrio, Eubacterium and Thermococcus. Quite a few genera are represented just by one species, e.g., Acetitomaculum, Acetonema, Chrysiogenes, Desulfobacula, Hippea, Stetteria and Succinispira. Autotrophic CO2-fixation is widespread among the acetogenic anaerobes such as Acetobacterium woodii, Clostridium aceticum and Moorella thermoautotrophica and especially among the methanogens of which only a few representatives are unable to grow with CO2 plus H2, e.g., Methanosaeta concilii, Methanosarcina acetivorans and the Methanosphaera species. A few sulfatereducing bacteria utilize CO2, such as Desulfobacterium autotrophicum and Desulfosarcina variabilis. The ability to fix molecular nitrogen is probably more common among anaerobes than known at the moment. Several clostridia are able to do so, with Clostridium pasteurianum being the first species demonstrated to have nitrogenase activity. Methanogens express active nitrogenase under nitrogen-limited growth conditions as has been demonstrated for Methanosarcina barkeri, Methanosarcina mazei and Methanococ-

e f iL

CHAPTER 1.4

The Anaerobic Way of Life

87

Table 1. Reactions yielding ATP by substrate-level phosphorylation in anaerobes. Reaction 1,3-Bisphosphoglycerate + ADP ¤ 3-phosphoglycerate + ATP Phosphoenolpyruvate + ADP ¤ pyruvate + ATP Acetyl phosphate + ADP ¤ acetate + ATP Butyryl phosphate + ADP ¤ butyrate + ATP Carbamoyl phosphate + ADP ¤ carbamate + ATP N10-Formyl FH4a + ADP + Pi ¤ formate + FH4 + ATP Glycine + 2H + ADP + Pi ¤ acetate + NH3 + ATP a

Enzyme

DGabs0 (kJ/mole)

Phosphoglycerate kinase Pyruvate kinase Acetate kinase Butyrate kinase Carbamate kinase Formyl-FH4 synthetase Glycine reductase

-24.1 -23.7 -12.9 -12.9 -7.5 +8.32 about -46.0

FH4, tetrahydrofolic acid.

cus maripaludis. Many more anaerobes can be expected to do so. So obligate anaerobes are known now for allimportant anaerobic habitats on earth. Because of their inability to utilize oxygen, they had to develop their strategies to conserve energy in the form of ATP, to metabolize substrates and to cope with some of their own products such as ethanol, lactate, butyrate or acetate. Some of the characteristic features of the anaerobes will be outlined.

Novel Ion Translocation Reactions Involved in Energy Conservation It is a fact that several anaerobic microorganisms produce ATP only by substrate-level phosphorylation. Growth on sugars or on amino acids coupled to the formation of ethanol, lactate, butyrate or acetate very often indicates that substrate-level phosphorylation is involved (Thauer et al., 1977). This holds true for lactic acid bacteria and also for many clostridia. Some of the reactions employed for ATP synthesis by these bacteria and by other anaerobes are listed in Table 1. It can be seen that the reactions 1 to 4 listed in Table 1 are part of the glycolytic pathway of acetate and butyrate formation. Carbamoyl phosphate is formed in the conversion of arginine to ornithine, and thereby becomes available for ATP synthesis. The conversion of N10formyl FH4 (N10-formyl tetrahydrofolic acid; an intermediate of methyl group oxidation) to formate, and FH4 gives rise to ATP synthesis. Glycine reductase is involved in the reductive part of the Stickland reaction, the pairwise fermenta-

tion of amino acids. This interesting reaction will be discussed in detail below. There are fermentations in which at first sight reactions giving rise to ATP synthesis cannot be identified. Such processes are for instance hydrogen-dependent fermentations; some are summarized in Table 2. Here it has been assumed for quite some time that electron transport processes might be coupled to ion translocation and that the ion-motive force generated might support ATP synthesis. Experimental proof for this assumption has been provided in recent years. Wolinella succinogenes grows on fumarate and H2 according to the equation given in Table 2. Clearly, this organism must gain ATP by electron transport phosphorylation. The electron transport chain that catalyzes this reaction (Fig. 1B) consists of hydrogenase, menaquinone and fumarate reductase (Lancaster and Kröger, 2000). Using vesicles and reconstituted liposomal systems the generation of a proton-motive force could be demonstrated in the course of H2dependent menaquione reduction as catalyzed by the hydrogenase (Gross et al., 1998). A number of other bacteria also can take advantage of ion-translocating electron transport system using fumarate as a terminal electron acceptor (Kröger et al., 1992). Formate, NADH or H2 are typical electron donors, and succinate or propionate are formed as catabolic end products. The pathway (as employed by the methanogens) for CO2-reduction to methane by H2 is depicted in Fig. 2. It has been demonstrated in recent years that one reaction, the methyl group transfer from methyltetrahydromethanopterin to coenzyme M, is coupled to the translocation of sodium ions (Deppenmeier et al., 1996). This

Table 2. H2-dependent fermentations. Reaction Fumarate + H2 CO2 + 4 H2 2 CO2 + 4 H2 SO42- + 4 H2 + H+ 2 FeOOH + H2 + 4 H+

Change of free energy Æ Æ Æ Æ Æ

succinate CH4 + 2 H2O CH3·COO- + H+ + 2 H2O HS- + 4 H2O 2 Fe2+ + 4 H2O

DGo¢ = -86kJ/mol DGo¢ = -131kJ/mol DGo¢ = -95kJ/mol DGo¢ = -152kJ/mol DGo¢ = -110kJ/mol

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CHAPTER 1.4 Succinate

Fumarate + 2 H+

A)

B) 2 H+

2 H+

Polysulfide reductase

heme b

2e

FAD

FeS

MKH2

_

MKb

heme b

Cytoplasm

di-heme b

MK FeS

Ni-Fe

2 H+ Hydrogenase H2

Fumarate reductase Ni-Fe

Mo

H+ + [S]

Periplasm

FeS

FeS

_

HS

H2

2 H+

Hydrogenase Fig. 1. Anaerobic respiration in Wolinella succinogenes. A) Polysulfide reduction: The membrane-bound hydrogenase is composed of three subunits (HydABC) and contains heme b, iron-sulfur clusters (FeS) and the nickel/iron center (Ni-Fe) for hydrogen oxidation. The gene products PsrA, B and C form the polysulfide reductase which contains a molybdopterin guanine dinucleotide (Mo), iron sulfur clusters (FeS). A menaquinone (Mkb) is tightly bound to the protein. Electron transfer is probably mediated by diffusion and collision of the enzymes. B) Fumarate reduction: The hydrogenase is identical to the one shown in Fig. 1A. The fumarate reductase consists of three subunits (frdCAB). A diheme cytochrome b anchors the enzyme in the membrane (di-heme b). The catalytic subunit carries a covalently bound FAD. These subunits are connected by an iron-sulfur protein (FeS). Electron transfer from the hydrogenase to the fumarate reductase is mediated by menaquinone.

system represents a novel type of sodium ion pump, which will be discussed below in connection with other sodium ion pumps. Some methanogens (e.g., Methanosarcina spp.) employ two novel membrane-bound electron transport systems generating an electrochemical proton gradient. The systems are composed of the heterodisulfide reductase and either a membrane-bound hydrogenase or an F420H2 dehydrogenase (Bäumer et al., 2000), which is functionally homologous to the protontranslocating NADH dehydrogenase (complex I of the respiratory chain). It has been shown that all of these enzymes are involved in proton translocation. Interestingly, the electron transport systems of these organisms contain electron carriers (such as cytochromes and the novel redox carrier methanophenazine), not found in methanogens utilizing only H2 + CO2 (Deppenmeier et al., 1999). A number of archaea as well as of bacteria reduce elemental sulfur with H2 to H2S (Hedderich et al., 1999). Examples are Pyrodictium occultum, Stetteria hydrogenophila and Desulfu-

robacterium thermolithotrophum, but also the already mentioned Wolinella succinogenes in which a H2:polysulfide reductase was characterized consisting of a nickel-iron hydrogenase, menaquinone and a molybdenum iron sulfidecontaining polysulfide reductase (Fig. 1A). Because the solubility of elemental sulfur in water is extremely low, it is believed that polysulfide is the actual electron acceptor (Hedderich et al., 1999). It is formed in an H2S environment according to: nS0 + HS - Æ S 2- n +1 + H +

(1)

Proton gradients are also established in the process of dissimilatory sulfate reduction. Here, the electron transfer from H2 to sulfite is coupled to ATP synthesis via a chemiosmotic mechanism (Badziong and Thauer, 1980). Shewanella putrefaciens (not an obligate anaerobe) can grow with Fe3+ and H2. The mode of energy conservation is not known as yet. Diffusion gradients may also be exploited for the generation of a proton-motive force. As long as the intercellular lactate concentration is high

CHAPTER 1.4 Fig. 2. Membrane-bound electron transport chain in Methanosarcina mazei. In the course of methanogenesis, methyl-coenzyme M (CH3-SCoM) is formed and is reductively cleaved by the methyl-CoM reductase which uses coenzyme B (HSCoB) as electron donor. The reaction results in the formation of methane and a heterodisulfide (CoB-S-SCoM) from HS-CoM and HS-CoB. The disulfide functions as electron acceptor of the anaerobic respiratory chain. Molecular hydrogen or reduced coenzyme F420 (F420H2) serves as electron donors. The F420H2 dehydrogenase contains FAD and FeS clusters and is responsible for the oxidation of F420H2. Electrons are transferred to methanophenazine (MPhen). The reduced form of this novel cofactor is the electron donor of the heterodisulfide reductase. This enzyme contains heme b and iron-sulfur clusters. It catalyzes the reduction of CoM-S-S-CoB. The H2dependent electron transport system is composed of a membrane-bound hydrogenase which is very similar to the corresponding enzyme from Wolinella (Fig. 1). Methanophenazine functions as mediator of electron transport to the heterodisulfide reductase.

The Anaerobic Way of Life in

cytoplasmic membrane

89

out 2 H+

Membrane-bound hydrogenase [NiFe]

2H+

MPhenH2

CoB-S-S-CoM + 2H+

CH3-S-CoM

2e– [FeS]

MPhen

Heterodisulfide reductase CH4

home b

H2

2e– [FeS]

home b

2 H+

CoM-SH + CoB-SH MPhenH2 MPhen

F420H2 dehydrogenase 2H+

methanogenic substrates

as compared to the extracellular one, it can be exported accompanied by two protons: Lactate inside + 2H + inside Æ lactate outside + 2H + outside (2) Thus, the proton/product symport helps lactate acid bacteria to increase their ATP yield (Konings et al., 1997).

Sodium Ion Pumps Cells have the tendency to expel sodium ions from the interior. Usually expulsion is catalyzed by sodium-proton antiporters, but a number of obligately anaerobic microorganisms have primary sodium ion pumps at their disposal. In these organisms certain exergonic reactions are coupled with Na+-translocation across the cytoplasmic membrane. One example was given already: the methyltetramethanopterin:coenzyme M methyltransferase reaction which is present in all methanogens and which is responsible for the Na+-dependence of growth and methane formation of this group of archaea. This

F420H2

[FeS] e–

F420

[FAD]

e– 2 H+

enzyme system is an extremely complex one consisting of eight different subunits and containing B12 as cofactor (Gottschalk and Thauer, 2001). A related enzyme system may occur in Acetobacterium woodii and related organisms that are Na+dependent and generate a sodium ion-motive force during acetogenesis (Heise et al., 1989). This, however, is not true for all acetogens. Organisms such as Clostridium aceticum and Moorella thermoautotrophica are not Na+ dependent; they contain cytochromes and apparently generate a proton gradient instead of a sodium ion gradient (Hugenholtz and Ljungdahl, 1990). Certain decarboxylases have been found to function as primary Na+ pumps. They are membrane bound and they contain biotin. These enzymes occur in organisms such as Propionigenium modestum, Acidaminococcus fermentans or Klebsiella pneumoniae, and the acids are decarboxylated with Na+ extrusion are oxaloacetate, methylmalonyl-CoA, glutaconyl-CoA or malonyl-acyl carrier protein (malonyl-ACP; Dimroth, 1997; Dimroth and Schink, 1998). A scheme is depicted in Fig. 3.

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CHAPTER 1.4 Fig. 3. Decarboxylation reactions coupled to sodium ion translocation: 1) Oxaloacetate decarboxylase (e.g., is used by Klebsiella pneumoniae to ferment citrate); 2) Methylmalonyl-coenzyme A (CoA) decarboxylase (e.g., is used by Propionigenium modestum for succinate metabolism); 3) Malonyl-S-acyl carrier protein (ACP) decarboxylase (e.g., is used by Malonomonas rubra growing on malonate); and 4) Glutaconyl-CoA decarboxylase (e.g., is used by Acidaminococcus fermentans to ferment glutamate).

2 Na+ out

H+

cytoplasmic membrane in

CO2 –

Biotin

Biotin-CO2

2 Na+

Citrate

Oxalacetate1

Pyruvate

Succinate

Methylmalonyl-CoA2

Propionyl-CoA

Propionate

Malonate

Malonyl-S-ACP3

Acetyl-S-ACP

Acetate

Glutamate

Glutaconyl-CoA4

Crotonyl-CoA

Butyrate

Degradative Pathways With respect to the degradation of substrates, the anaerobes have disadvantages and advantages. One difficulty is that in the absence of an external electron acceptor anaerobes must balance their oxidation and reduction reactions. The electron donors and acceptors are derived from organic molecules of medium redox states such as sugars, organic acids, heterocyclic compounds and amino acids. Often more reduced (e.g., ethanol) and more oxidized (e.g., CO2) products are formed. In a few fermentations, the redox state of the substrate and the product is the same, e.g., the fermentation of hexoses to two lactates or three acetates. Highly oxidized or reduced compounds such as carbon dioxide or hydrocarbons, respectively, are only suitable for fermentation together with inorganic electron donors or acceptors. Another disadvantage of anaerobes is, of course, that oxygen cannot be employed for the initial attack of certain substrates such as hydrocarbons. On the other hand, there are a number of advantages. Oxygen-sensitive systems can be taken advantage of radical reactions or even of radical enzymes. So under the dictate of balanced redox reactions and with the involvement of unique enzymes and reactions, a fascinating array of unusual fermentations has evolved; some will be discussed now.

Coenzyme B12-Dependent Pathways When Clostridium tetanomorphum or Clostridium cochlearium grows on L-glutamate, the

NH2 COOH HOOC L-glutamate

0.2 acetate, 0.4 butyrate, 1 CO2, 0.2 H2

1 NH2 COOH HOOC CH3

CH3-CO-COOH pyruvate

L-threo-β-methylaspartate 2

CH3–COOH

NH2

4

COOH

3

HOOC CH3 mesaconate

COOH HOOC

H2O

1 acetate

CH3

OH citramalate

Fig. 4. Pathway of L-glutamate fermentation by Clostridium tetanomorphum: 1) Glutamate mutase (coenzyme B12dependent); 2) b-Methylaspartase; 3) Citramalate dehydratase; and 4) Citramalate lyase.

substrate is prepared for a cleavage into a twocarbon and a three-carbon compound in an interesting way. Under the catalysis of glutamate mutase (a B12-containing enzyme), L-glutamate is converted to L-threo-b-methylaspartate (Buckel and Golding, 1996b). This carbonskeleton rearrangement facilitates the elimination of ammonia and formation of mesaconate by b-methylaspartase. Subsequently, mesaconate is hydrated to citramalate, which then is cleaved into acetate and pyruvate (Buckel, 1980; Fig. 4). Oxidative decarboxylation of pyruvate results in

CHAPTER 1.4

The Anaerobic Way of Life

Fig. 5. Interconversion of succinate and propionate by methylmalonylCoA mutase: 1) Propionate CoAtransferase; 2) Methylmalonyl-CoA mutase (coenzyme B12-dependent); 3) Methylmalonyl-CoA epimerase; and 4) Transcarboxylase (biotincontaining).

O

1

COOH

HOOC succinate

HOOC

O CH3 AoCS propionyl-CoA

91

SCoA succinyl-CoA

CH3

HOOC

propionate

O

2

COOH HOOC oxaloacetate 4

CH3H

O

CH3 H

SCoA HOOC

CH3 HOOC pyruvate

O (s)-methylmalonyl-CoA

SCoA

HOOC

3

O (R)-methylmalonyl-CoA

gluconeogenesis

HO

ATP

O

2 HO

dihydroxyacetone

1

OH OH

glycerol H2O

NADH + H+

3 HO

O

3-hydroxypropion- 4 aldehyde

HO

O O P O– O– dihydroxyacetone phosphate

ADP HO

OH

O

glyceraldehyde3-phosphate

catabolism OH

1,3-propanediol

Fig. 6. Pathway of glycerol fermentation by Citrobacter freundii: 1) Glycerol dehydrogenase; 2) Dihydroxyacetone kinase; 3) Glycerol dehydratase (coenzyme B12-dependent); and 4) 1,3-Propanediol dehydrogenase.

the formation of acetyl-CoA and reduced ferredoxin, which is reoxidized during the synthesis of butyryl-CoA from two moles of acetyl-CoA. Then, ATP is synthesized in the acetate and butyrate kinase reactions (Barker, 1981). By this pathway, a degradation of glutamate via the tricarboxylic acid cycle is circumvented; the latter would not be feasible because of an unbalanced generation of reducing equivalents in the form of NADH and FADH2. Coenzyme B12-dependent rearrangements like the glutamate mutase reaction proceed via radical intermediates; they are per se oxygen sensitive although another reaction of this type, the methylmalonyl-CoA mutase reaction, proceeds in higher eukaryotes such as man. This reaction is also of key importance in propionic acid bacteria and many other anaerobes because

it allows the interconversion of succinate and propionate (Fig. 5). A fermentation that involves a coenzyme B12dependent reaction and proceeds only under anaerobic conditions is the glycerol conversion to 1,3-propanediol. This fermentation was discovered in enteric bacteria such as Citrobacter freundii and Klebsiella pneumoniae; it proceeds as depicted in Fig. 6. Glycerol is oxidized to dihydroxyacetone, which is converted further to dihydroxyacetone phosphate. To balance the fermentation, a portion of glycerol is dehydrated to 3-hydroxypropionaldehyde in a coenzyme B12dependent reaction. Subsequently, the aldehyde is reduced to the major fermentation product 1,3-propanediol, which is of great biotechnological interest. The bottleneck of the pathway is the coenzyme B12-dependent glycerol dehydratase

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CHAPTER 1.4

that is rapidly inactivated during glycerol dehydration (Daniel et al., 1998).

Degradation of Amino Acids and a-Hydroxy Carboxylic Acids Novel reactions occur in a number of anaerobes for the utilization of a-amino acids and ahydroxy carboxylic acids. If redox balance allows, these acids can be oxidized, of course, to the corresponding a-keto acids and then very easily metabolized further. So lactate or alanine can be oxidized to pyruvate and further to acetyl-CoA. This often is not possible because an acceptor for the electrons generated is not available. A commonly used pathway involves the reduction of the a-keto acids generated by deamination of amino acids to the corresponding hydroxy carboxylic acids, followed by activation to the CoA ester and dehydration to an enoyl-CoA (Fig. 7). A simple dehydration of ahydroxy carboxylic acids is not feasible because it would have to proceed against the rule of Markovnikov (Jones, 1961). A well-studied example is the dehydration of a-hydroxyglutaryl-CoA to glutaconyl-CoA carried out by Acidaminococcus fermentans. The enzyme, a-hydroxyglutarylCoA dehydratase, is extremely oxygen sensitive and contains [Fe-S] clusters, reduced riboflavin, and FMNH2. The activation of the dehydratase is catalyzed by an activator protein and requires a reducing agent and catalytic amounts of ATP and Mg2+. A novel mechanism involving thiol ester-derived radical anions (ketyls) has been postulated for these dehydrations (Buckel, 1996a).

NADH + H+ NAD+ COOH

H R

O α-keto acid

1

H COOH

R

OH α-hydroxy acid acetyl-CoA 2

H2O

H SCoA

enoyl-CoA

3

1 Alanine Æ 1 acetate + 1 CO 2 + 1 ammonia + 4H 2 Glycine + 4H Æ 2 acetate + 2 ammonia The structurally related compounds betaine and sarcosine can also serve as hydrogen acceptors (Naumann et al., 1983; Hormann and Andreesen, 1989), methylamines being formed instead of ammonia: Betaine + 2H Æ trimethyla min e + acetate Sarcosine + 2H Æ methyla min e + acetate Acetate formation from glycine proceeds via acetyl phosphate, and the last step of acetate formation is catalyzed by acetate kinase giving rise to ATP synthesis by substrate-level phosphorylation. The key enzyme of glycine fermentation (glycine reductase) was well studied in Eubacterium acidaminophilum (Andreesen, 1994). The enzyme consists of four proteins including one selenoprotein (enzyme A), a pyruvoyl-protein (enzyme B), enzyme C, and thioredoxin. The reaction mechanism is depicted in Fig. 8. The pyruvoyl residue of enzyme B forms a Schiff-base with glycine, which then reacts with the Se--anion of protein A to yield a carboxymethylselenocysteine residue linked to protein A and the iminopyruvoyl protein. Subsequently, ammonia is released by hydrolysis or in the next turnover. Elimination of ketene yields the oxidized protein A-Se-S intermediate, which is reduced by thioredoxin. Reduction of thioredoxin is catalyzed by thioredoxin reductase with NADH or another electron donor. The hypothetical ketene intermediate adds to the cysteine residue of protein C. An acetylcysteine is formed, which is cleaved by phosphate (Pi) to form acetyl phosphate. Again, this is a complex reaction, which only can be visualized to occur in anaerobes.

acetate

O R

Another way to deal with certain a-amino acids is reductive deamination. Such deaminations are part of the Stickland reaction in which amino acids are fermented pairwise. Alanine, for instance, is oxidized and the reducing equivalents generated are transferred to glycine:

O

SCoA OH α-hydroxyacyl-CoA R

Fig. 7. a-Hydroxy acid pathway: 1) a-Hydroxy acid dehydrogenase; 2) CoA transferase; and 3) a-Hydroxyacyl-CoA dehydratase.

Degradation of Aromatic Compounds and Hydrocarbons Most of the aromatic compounds studied to date are first transformed to benzoyl-CoA, the central intermediate of the best-studied pathway for anaerobic degradation of aromatic compounds (Harwood et al., 1999). Benzoyl-CoA then

CHAPTER 1.4 Fig. 8. Mechanism reductase.

The Anaerobic Way of Life of

glycine

H2O

93

O

B

CH3

N O

+H N 3

H

COOH glycine

pyruvoyl-enzyme B

O B

CH3

N HN+

H

NH4+

O

B

CH3

N

COOH

NH2+

H –Se

selenoprotein A

HOOC Se

A

A

HS HS

NAD+ Se

thioredoxin

A

NADH

S [H2C

C O] ketene

HS

C

enzyme C

Pi

O

O C

H3C

OPO32–

H3C

S

acetyl phosphate

undergoes a reductive attack (Schink et al., 2000). The key enzyme for this attack is the benzoyl-CoA reductase, which was purified from the denitrifying bacterium Thauera aromatica and characterized as a FAD- and iron-sulfur cluster-containing enzyme complex (Boll and Fuchs, 1995). Under hydrolysis of ATP, one electron is added to the thiol ester carbonyl of benzoyl-CoA and the resulting radical intermediate is reduced further to cyclohexa-1,5-dienecarboxyl-CoA (Buckel and Golding, 1999; Fig. 9). This reaction may be of general importance for the anaerobic degradation of aromatic compounds. Recently, it was shown that the reductive strategy for destabilization of the ring is not the only one used in anaerobic degradation of aromatic compounds. Anaerobic degradation of 3,5-dihydroxybenzoate by Thauera aromatica (Philipp and Schink, 2000) and 1,3-dihydroxybenzene by Azoarcus anaerobius (Philipp and Schink, 1998) proceeds by a novel mechanism. Phenolic compounds with their hydroxyl groups in meta position to each

other are hydroxylated by membrane-bound enzymes yielding hydroxyhydroquinone, which is later dehydrogenated to the nonaromatic compound hydroxybenzoquinone. Thus, oxidation rather than reduction is used to overcome the stability of the aromatic ring.

Radical Enzymes Glycyl radical enzymes are involved in a number of anaerobic reactions. Well-studied examples are the pyruvate formate lyase (Knappe et al., 1984), the anaerobic ribonucleotide reductase (Licht et al., 1996), and the benzyl succinate synthase (Leuthner et al., 1998). The latter initiates the breakdown of toluene under anaerobic conditions. These glycyl radical enzymes are formed from their precursor enzyme in a reaction, which requires S-adenosyl methionine. The pyruvate formate lyase of Escherichia coli is synthesized

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CoAS

CHAPTER 1.4 O

CoAS

O

H+ , e

H+ , e

2 ATP + 2 H2O

O

CoAS

H

2 ADP + 2 Pi

benzoyl-CoA H

H cyclohexa-1, 5-diene-carboxyl-CoA

Fig. 9. Mechanism of benzoyl-CoA reductase.

as an inactive and coenzyme-free protein. The enzyme is posttranslationally modified by Sadenosyl methionine and a reduced flavodoxin in a reaction catalyzed by an activase. A hydrogen atom is abstracted from a specific glycine residue, yielding methionine and 5¢-deoxyadenosine from S-adenosyl methionine. The formed free radical (HS-enzyme) is involved in a twostep reaction: Pyruvate + HS-enzyme Æ acetyl - S-enzyme + formate Acetyl- S-enzyme + CoASH Æ acetyl - SCoA + HS-enzyme Pyruvate formate lyase, like the other glycyl radical enzymes, is rapidly inactivated by oxygen. Anaerobic alkane-degrading bacteria have also been isolated recently. Alkanes are used as substrates by several species of sulfate-reducing microorganisms (Aeckersberg et al., 1998). Another group of anaerobic hydrocarbondegrading bacteria is dependent on syntrophic associations with methanogens. The biochemistry of the process is still poorly understood but it can be speculated that again radicals are generated to initiate this breakdown (Zengler et al., 1999). A number of potentially hazardous compounds in our environment are halogenated (e.g., pentachlorophenol or perchloroethene). These compounds can be partially or completely degraded under anaerobic conditions. This degradation occurs by reductive dehalogenations. Organisms such as Desfulfitobacterium dehalogenans, Dehalobacter restrictus or Dehalospirillum multivorans contain corrinoid-proteins, which exhibit dehalogenase activities (Holliger et al., 1999). There is evidence that these H2-dependent fermentations are also coupled with the generation of a proton-motive force.

Anaerobic Food Chains The anaerobic degradation of complex organic matter depends on the cooperation of various

trophic groups of anaerobic bacteria and archaea. Two possible schemes for anaerobic food chains, as they occur in nature in the absence or in the presence of sulfate, are presented in Fig. 10. Polymers such as polysaccharides, proteins and nucleic acids are initially converted to oligomers and monomers and subsequently fermented by the “classical” primary fermentative bacteria. In the absence of sulfate, the products acetate, methanol, methylamines, CO2 and H2 can be used directly by methanogenic bacteria to convert them to methane and carbon dioxide. Alcohols longer than one carbon atom, fatty acids longer than two carbon atoms and branched or aromatic fatty acids are degraded by the secondary fermenters to acetate, C1-compounds and H2, which are subsequently used by the methanogens. Because the reactions catalyzed by the secondary fermentative bacteria are mostly endergonic under standard conditions, they depend on a very efficient cooperation with the subsequent partners. Such cooperations are called syntrophic relationships, in which the pool size of shuffling intermediate has to be kept small to allow efficient degradation. In sulfate-rich anaerobic habitats, such as marine sediments, sulfate-reducing bacteria further degrade the primary fermentation products. As many sulfate reducers are metabolically more versatile than methanogenic bacteria, they can use and oxidize all classical fermentation products to carbon dioxide, simultaneously reducing sulfate to sulfide (Hansen, 1994; Jansen and Hansen, 1998; Zengler et al., 1999; Fig. 10B). In addition to the primary fermentations that have already been mentioned, three important points should be briefly discussed here: the fate of acetate under anaerobic conditions, production of H2, and the syntrophic relationships. Acetate is the end product of a number of fermentations starting from substrates with two (e.g., ethanol) or more carbon atoms (e.g., glucose), but it is also produced by acetogenic organisms from one-carbon compounds (e.g.,

CHAPTER 1.4

The Anaerobic Way of Life

methanol) and from H2 + CO2. Because so many pathways lead to the formation of acetate under anaerobic conditions, the further degradation of acetate is of great importance for carbon flow under anaerobic conditions. Among the methanogenic archaea, only species of the genera Methanosarcina, Methanosaeta and Methanothrix are able to utilize and degrade acetate to methane and carbon dioxide (e.g., Methanosarcina barkeri, Methanothrix thermophila and Methanosaeta concilii). The degradation occurs according to the following equation (Thauer et al., 1989):

polymers (proteins, polysaccharides, lipids, nucleic acids)

monomers and oligomers (peptides, amino acids, sugars, acids purins, pyrimidines, glycerol)

alcohols, propionate, butyrate, lactate, other products

CO2+ H2

CH 3-COOH Æ CH 4 + CO 2 DG 0¢ = -36 kJ mol

acetate

methanol, methylamines

formate

methanogenesis

polymers (proteins, polysaccharides, lipids, nucleic acids)

B

monomers and oligomers (peptides, amino acids, sugars, acids, purins, pyrimidines, glycerol)

alcohols, propionate, butyrate, lactate, other products

acetate + H2

CO2

95

sulfidogenesis

Initially acetate is activated to acetyl-CoA by acetate kinase and phosphotransacetylase or directly by acetyl-CoA synthetase (Methanosaeta). Acetyl-CoA is subsequently bound to the carbon monoxide (CO) dehydrogenase complex, at which it is decarbonylated by cleavage of the carbon-carbon bond. The methyl-group is subsequently transferred via tetrahydromethanopterin (THMP) to coenzyme M, and CO is oxidized to CO2, providing the reducing equivalents for the reduction of the methyl-coenzyme M to methane by the pathway shown in Fig. 11 (Thauer, 1998; Ferry, 1997; Ferry, 1999). It is interesting that the CO dehydrogenase complex, which catalyzes the decarbonylation of acetylCoA to methyl-THMP and CO and the oxidation of CO, also catalyzes the reactions mentioned in a reversible manner. In methanogens utilizing acetate, the direction of decarbonylation predominates; when, however, organisms such as Methanobacterium thermoautotrophicum grow with H2 + CO2, they use this enzyme system to synthesize acetyl-CoA from methyl-coenzyme M and CO for autotrophic growth (Zeikus, 1983; Fuchs, 1986; Shieh and Whitman, 1988; Huber and Wachtershäuser, 1997). Similarly, acetogenic bacteria such as Acetobacterium woodii and Moorella thermoacetica produce acetyl-CoA from methyl-tetrahydrofolate and CO (Wood et al., 1986; Ljungdahl, 1986; Shanmugasundaram et al., 1988; Menon and Ragsdale, 1999). A number of sulfate-reducing bacteria are also able to oxidize acetate completely to CO2 under anaerobic conditions: 2-

CH 3-COOH + SO4 + H + Æ 2CO 2 + HS - + 2H 2O Fig. 10. Anaerobic food chains. (A) Methanogenesis. As a terminal process, all organic material is metabolized to methane via a few methanogenic substrates: CO2 + H2, acetate, formate, methanol and methylamines. (B) Sulfidogenesis. As a terminal process, incomplete oxidizers convert various products to CO2 and acetate, and the complete oxidizers couple sulfate reduction with acetate oxidation to CO2. In addition, H2 can be used for sulfate reduction.

Most of them also take advantage of the described C1-pathway with the CO dehydrogenase complex for decarbonylating acetyl-CoA. The pathway is investigated in more detail in Desulfotomaculum acetoxidans, Desulfobacterium autotrophicum and in the archaeon Archaeoglobus fulgidus (Spormann and Thauer,

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CHAPTER 1.4

2) The NADH-ferredoxin oxidoreductase:

CH3 -COO– ATP

HS-CoA

ADP

CH3 -CO-S-CoA

THMP

CH3 -THMP

CO HS-CoM

CH3 -S-CoM H2O

CoB-SH 2H CoB-S-S-CoM

CO2

CH4

Fig. 11. Conversion of acetyl-CoA to methane and carbon dioxide. THMP, tetrahydromethanopterin; HS-CoM, coenzyme M.

1988; Hansen, 1994; Möller-Zinkhahn et al., 1989; Brüggemann et al., 2000). Only a small number of acetate oxidizing, sulfate reducers (e.g., Desulfobacter postgatei) use the tricarboxylic acid cycle to carry out acetyl-CoA oxidation (Brandis-Heep, 1983; Möller et al., 1987; Thauer, 1988; Thauer et al., 1989). Activation of acetate in D. postgatei occurs by a succinyl-CoA:acetate CoA-transferase; acetate kinase and phosphotransacetylase are lacking. Many fermentation reactions are associated with the evolution of molecular hydrogen, H2. This allows a shift from producing alcohols and lactate to acetate and butyrate, a shift beneficial to the organisms because the ATP yield is increased. Important precursors of H2 are formate and reduced ferredoxin, and H2 formation is catalyzed by formate hydrogenlyase and hydrogenase, respectively. There are two important reactions coupled to ferredoxin reduction and ultimately to H2 formation: 1) The pyruvate-ferredoxin reaction:

oxidoreductase

Pyruvate + Fd ox ¤ Fd red + acetyl -CoA + CO 2 DG 0 ¢ = -19.2kJ mol The reaction is exergonic so that it can drive H2 formation even at a hydrogen partial pressure (PH2) of 1.013 kPa. The enzyme was first purified from Clostridium acidiurici (Uyeda and Rabinowitz, 1971; Charon et al., 1999).

NADH + Fd ox ¤ Fd red + NAD + + H + DG 0¢ = +18.8kJ mol This reaction was discovered in C. kluyveri (Jungermann et al., 1969; Gottschalk and Chowdhury, 1969); it is endergonic and will only proceed at a largely reduced PH2. In anaerobic habitats, the PH2 is kept as low as 10 Pa by H2consuming organisms such as the methanogenic archaea, and acetogenic and sulfidogenic bacteria. Hydrogen consumption by these microorganisms results in the phenomenon of interspecies hydrogen transfer, which has two consequences. First, the product patterns of saccharolytic fermentations as carried out by many clostridia are changed; for example, glucose can be fermented to acetate and CO2. The second consequence of the generation of a low PH2 by the hydrogen-consuming bacteria is that it opens up an ecological niche for a fascinating group of anaerobes, the obligate proton-reducing bacteria. These organisms, were first described in 1967, when a culture called “Methanobacillus omelianskii” was found to consist of two different organisms carrying out two different fermentations (Bryant et al., 1967): 1) The “S” organism carries out ethanol oxidation: CH 3-CH 2-OH + H 2O Æ CH 3-COOH + 2H 2 2) A methanogenic archaeon consumes molecular hydrogen for methane production: 2H 2 + 1 2 CO 2 Æ 1 2 CH 4 + H 2O Cocultures of this type were termed “syntrophic” cultures, because the organisms involved mutually depend on one another. Molecular H2 evolution allows fermentative growth of the “S” organism, but only if the PH2 is kept low enough by the methanogenic bacterium. The term “interspecies hydrogen transfer” was coined for this kind of connection between H2 evolution and H2 consumption. Other examples for syntrophically ethanol-oxidizing bacteria known today are Thermoanaerobacterium brockii (Ben-Bassat et al., 1981), Pelobacter species (Schink, 1984; Schink, 1985), and in the absence of sulfate, Desulfovibrio vulgaris (Bryant et al., 1977). Not only alcohols but also organic acids can be oxidized to acetate and H2 this way, such as propionate by Syntrophobacter pfennigii (Wallrabenstein et al., 1995) and butyrate by Syntrophomonas species (Roy et al., 1986; McInerney et al., 1981). As these oxidations are more endergonic than alcohol oxidations, PH2 has to be decreased to significantly lower values ( Thr > Ser > Trp > (Asp, Glu, Arg).

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CHAPTER 1.7

Table 3. Degradative chemical reactions and isomerization reactions important to irreversible protein denaturation, especially at elevated temperature. Reaction Deamidation Racemization Isomerization Glycation Oxidation

Proteolysis Photodegradation

Amino acids involved Asn, Gln (especially in Asn-Gly and Asn-Ser sequences) Asp Pro (cis-trans isomerization) Lys and other amino acids reacting with reducing sugars Cys ¤ sulfenic Æ cysteic acid (oxidation or SH/SS exchange via mixed disulfides) Met ¤ sulfoxide Æ sulfone Polypeptides Æ amino acids Trp Æ kynurenin Æ N-formyl kynurenin Tyr Æ DOPA, dityrosine Cystine Æ 2Cys

Comments Independent of pH, product: iso-Asp as substrate of methyl transferase, leading to repair or clearance. Catalyzed by peptidyl prolyl-cis-trans isomerases.a Cross-linking by Maillard reactions, involved in in vivo degradation.b Thiolate mechanism catalyzed by Cu2+ or Fe2+ or protein disulfide isomerases (PDI, DsbA/DsbB, etc.). Significant both in vivo and in vitro in the presence of oxygen radicals. Caused either by proteases or autolysis, or by H+-catalyzed peptide cleavage; nonenzymatic, between Asp and Pro and Asp and (C-terminal) Asn. Caused by nonionizing or ionizing radiation, depending on the local microenvironment of the amino acids.

Abbreviations: Asn, asparagine; Gln, glutamine; Ser, serine; Asp, aspartate; Lys, lysine; Pro, proline; Cys, cysteine; Met, methionine; PDI, protein disulfide isomerase; DsbA/DsbB, disulfide-bond forming proteins; and DOPA, dihydroxyphenylalanine. a Schiene-Fischer and Fischer (2000). b cf. Barrett (1985). For further references and details, cf. Greenstein and Winitz (1961); Meister (1965); Cecil (1963); Gottschalk (1972); Freedman (1973); Barrett (1985); Stadtman (1990); Stadtman and Oliver (1991); Volkin et al. (1995); Berlett and Stadtman (1997); Jaenicke and Seckler (1997); Daniel and Cowan (2000); Jaenicke and Lilie (2000); Schiene-Fischer and Yu (2001); and Vieille and Zeikus (2001).

Cysteine exhibits low stability: Depending on temperature and pH, it undergoes either oxidation (to form cystine), or elimination of sulfur (lanthionin formation). The lower limit at which degradation in aqueous buffer solutions was detectable was ca. 110∞C (Bernhardt et al., 1984). One may assume that up to this temperature range, biosynthesis can still balance the thermal decomposition. In the temperature regime of hydrothermal vents, e.g., at 250∞C (265 bar), the half-lives of the amino acids, peptides and proteins undergoing degradation were found to be too short to be offset by biosynthesis of these molecules (White, 1984). ATP and ADP hydrolysis become significant between 110 and 140∞C (Leibrock et al., 1995). This upper temperature limit coincides with the temperature range at which the hydrophobic hydration of nonpolar residues in aqueous solution vanishes (Sturtevant, 1977; Privalov, 1979; Baldwin, 1986; Jaenicke, 1991b; Jaenicke, 2000a). In summarizing the biochemical limitations of viability from the point of view of water-soluble proteins, temperatures beyond ca. 130–140∞C are not tolerable, for two reasons: 1) natural amino acids are hydrothermally decomposed and 2) the solvent properties of water are altered, blurring the difference between polar and nonpolar residues, thus interfering with the “hydrophobic collapse” (as the initial step of protein folding) and the formation of the densely packed hydropho-

bic core (as the prerequisite of protein stability). For nucleoproteins and lipoproteins or membranes, the same holds true because of the temperature limits of the intermolecular interactions between the polar and nonpolar components in the respective complexes. For both classes of proteins, extrinsic factors and compatible solutes may enhance the stability as well as the limits of growth (see below). The given upper temperature limit of viability has been confirmed for cells of the most extreme hyperthermophiles Pyrolobus fumarii, and strain 121 with its temperature of maximal growth at 121∞C (Blöchl et al., 1997; Kashefi and Lovley, 2003). Whether the protective action of compatible solutes and/or crowding induced by high levels of molecular chaperones contribute to this extreme thermotolerance, needs further investigation (cf. Carpenter et al., 1993; Zimmerman and Minton, 1993; Somero, 1995; Trent et al., 1997; Minton, 2000). Clearly, the biochemical limit of viability depends not only on the intact organization of the cell’s standard high-molecular weight components, but also on the low-molecular weight compounds such as coenzymes and metabolites. Again, in general, extremophiles make use of the common repertoire of compounds known from the metabolism of mesophiles. Keeping in mind the high catalytic rate of most enzymes under physiological conditions, the majority of meta-

CHAPTER 1.7

bolites do not limit viability at temperatures close to 100∞C. The reported half-lives of ATP and ADP range from ~1–6 hours at 100∞C, depending on the pH and the presence of metal ions (Ramirez et al., 1980; Leibrock et al., 1995; Daniel et al., 1996). However, the oxidized nicotinamide adenine dinucleotide (NAD+) has a half-life at 100∞C of no more than 10 min. To cope with this instability, nature can make use of at least four strategies: 1) high catalytic turnover, or 2) channeling of labile intermediates, 3) local stabilization in enzyme-ligand complexes, and 4) usage of an alternate metabolic pathway or a different, more stable compound. In the case of 3), the high affinity of ligands for their respective enzymes has frequently been shown to cause mutual stabilization (Danson, 1988; Jaenicke et al., 1996; Dams and Jaenicke, 1999).

Adaptive Stabilization Mechanisms of Nucleic Acids The integrity of nucleic acids is threatened at high temperatures, which can induce either strand separation and chemical damage of the nucleotide constituents or, at the extreme, breakage of backbone phosphodiester bonds (Grogan, 1998; Daniel and Cowan, 2000).

Mechanisms to Avoid Strand Separation An increased G+C content is known to increase the temperature Tm at which melting, i.e., strand separation of DNA and RNA occurs. Thus, a possible adaptation mechanism of nucleic acids to thermophilic and even more to hyperthermophilic conditions would be an increase in G+C. Indeed, a systematic study revealed a strong positive correlation between the G+C content of tRNAs and rRNAs with the optimum growth temperatures of prokaryotes (Galtier and Lobry, 1997; Fig. 6A). The same study showed, however, that the G+C content of genomic DNA is not correlated with the growth temperature (Fig. 6B). Quite the contrary, the DNA of some of the most hyperthermophilic archaea has a strikingly low G+C content, with values as low as 31 mol%, e.g., for Acidianus fervidus and Methanococcus igneus (Tmax >90∞C), and an average of ca. 45 mol% for all presently known hyperthermophilic archaea and bacteria (Stetter, 1996; Grogan, 1998). These data clearly suggest that in these organisms, the DNA double helix must be stabilized either by extrinsic factors such as ions and small metabolites or by proteins. It has been known for a long time that the addition of salts or polyamines leads to an increase in Tm. Actually some, but not all, hyperthermophiles accumulate high concentrations of putative

Life at High Temperatures

185

ionic thermoprotectants such as potassium diinositol-1¢,1¢-phosphate and tripotassium cyclic2,3-diphosphoglycerate (Hensel and König, 1988; Scholz et al., 1992). However, there is no clear correlation between the level of polycationic polyamines and growth temperature (Kneifel et al., 1986). In a number of archaeal hyperthermophiles, two unrelated groups of highly basic proteins were identified, which bind to DNA without marked sequence preference. Both the members of the HMf histone family, which are homologs of the eukaryal core histones, and the histone-like proteins from Sulfolobus species, for which no eukaryal homologues are known, increase the Tm of the DNA double helix significantly (McAfee et al., 1996; Soares et al., 1998). Thus, there is clear evidence that hyperthermophiles make use of different strategies to prevent DNA strand separation at their extreme growth temperatures. Certainly, the physiological interpretation of in vitro Tm data gained from topologically open molecules has to be taken with a grain of salt because cellular DNA is in a topologically closed conformation, and denaturation will not result in two independent single-stranded molecules, but in a randomcoil structure with intertwined strands (Marguet and Forterre, 2001). As a result, topologically closed DNA is undoubtedly more resistant to denaturation than open DNA. It was postulated that the introduction of positive supercoils into closed DNA, which is catalyzed by reverse gyrases from hyperthermophiles, specifically stabilizes the double helix and keeps it in a functional state at high temperature (Forterre et al., 1996; Lopez-Garcia and Forterre, 1997, 2000). However, the hyperthermophile Thermotoga maritima contains both “normal” and reverse gyrases and propagates negatively supercoiled plasmid DNA (Guipaud et al., 1997). tRNA molecules are not permanently integrated into larger macromolecular complexes. Therefore, in adapting to high temperatures, they must have developed mechanisms for intrinsic stabilization. Part of the stabilization energy may originate from an increased G+C content. However, unfractionated tRNA from the hyperthermophiles Pyrococcus furiosus and Pyrodictium occultum showed Tm values around 100∞C, too high to be attributable to the measured G+C content (Kowalak et al., 1994). An early investigation identified a broad variety of covalent posttranscriptional modifications in nucleosides from tRNA preparations of thermophiles and hyperthermophiles, six of which were structurally novel in showing alterations of their bases as well as methylation of their ribose moiety (Edmonds et al., 1991). Altogether, 23 modified nucleosides were identified in Pyrococcus furiosus; three of them (Fig. 7) not only

CHAPTER 1.7

80

70

60

tRNA G+C content %

n=224 genera 70 60 50 40 30 20 10 20

n=51 genera

23S rRNA stems G+C content %

80

B

n=165 genera

Genomic G+C content %

A

R. Jaenicke and R. Sterner 16S rRNA stems G+C content %

186

30 40 50 60

70

80 90 100 110

Optimal growth temperature °C

70 65 60 55

n=38 genera 80

70

60

5S RNA G+C content %

n=71 genera 70

60

50

40 10 20

30 40 50 60

70

80 90 100 110

Optimal growth temperature °C Fig. 6. G+C contents of A) various RNAs and B) genomic DNAs plotted against optimal growth temperatures. Data taken from Galtier and Lobry (1997).

N

HN CH3 N CH3

N

HN—COCH3

O

O

O

N

HO

CH3

HN S

O

HN

N

O

OCH3

N

HO

HO HO

O

HO OH

N2,N2,2’-O-trimethylguanosine (m22Gm) 5-methyl-2-thiouridine (m5s2U)

HO

OCH3

N -acetyl-2’-O-methylcytidine (ac4Cm) 4

Fig. 7. Modified nucleosides implicated in the stabilization of hyperthermophile tRNA. From Daniel and Cowan (2000).

CHAPTER 1.7

exhibited enhanced relative abundance with increasing growth temperature, but also higher stability, which they effected by 1) restricting the conformational flexibility of the ribose ring, 2) favoring the A-type helix, and 3) preventing phosphodiester-bond hydrolysis (Inoue et al., 1987; Kawai et al., 1992; Kowalak et al., 1994; Cummins et al., 1995). Apparently, the protecting effect of posttranscriptional tRNA modification is not restricted to the archaea: both the level of 5-methyl-2thiouridine and the Tm value of tRNA from the bacterium Thermus thermophilus show a significant increase with increasing growth temperature (Watanabe et al., 1976). The effect becomes even more compelling if tRNAs from psychrophiles are included in the comparison. While the abundance and the variety of posttranscriptional tRNA modifications are more pronounced in thermophiles and hyperthermophiles than in mesophiles, significantly less modifications are found in tRNAs from psychrophiles (Dalluge et al., 1997). The most abundant one is dihydrouridine, whose nonplanar base resists stacking, this way decreasing stability. In addition, dihydrouridine favors the C-2¢-endo sugar conformation, which is less rigid than the C-3¢-endo conformer (Yokoyama et al., 1981). Obviously, enhanced flexibility is essential for optimal functioning at low temperature, whereas high intrinsic stability has lower priority. In the case of rRNAs, significant stabilization is provided by their conjugation with proteins within the ribosomal complex. In accordance with this argument, the levels of posttranscriptional modifications of rRNAs are much lower than in tRNAs, both in mesophiles and in thermophiles. Still, rRNA modifications are much more abundant in Sulfolobus solfataricus than in Escherichia coli, and the level of stabilizing ribose O-2¢ methylations significantly increases with the culture temperature of the hyperthermophile (Noon et al., 1998).

Mechanisms to Avoid and Repair Chemical Damage of Nucleotides Chemical damage of nucleic acids by hydrolytic attack close to the boiling point of water is an enormous potential threat for hyperthermophiles. The most common damages to DNA are 1) base deamination, 2) loss of bases from one strand with apurinic or apyrimidinic sites as final products, and 3) hydrolytic cleavage of phosphodiester bonds. It was suggested that, above 100∞C, DNA would be subject to a ca. 3000-fold increase in the levels of deamination and depurination compared with DNA at 37∞C (Lindahl, 1993). Furthermore, it was estimated from in vitro stability data that under the physiological conditions

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of S. solfataricus (intracellular pH 6, 80∞C) two apurinic sites per gene per cell division would accumulate (Grogan, 1998). The most severe damage of nucleic acids is the hydrolytic cleavage of the backbone phosphodiester bond. For this reaction, it was postulated that the preceding depurination at an adjacent site is the ratelimiting step (Marguet and Forterre, 1998; Marguet and Forterre, 2001). In contrast, for RNA, hydrolytic strand breakage is not coupled to depurination; instead it occurs via the direct attack of the phosphodiester bond by the ribose 2¢-OH oxygen. In vitro, at around 100∞C, singlestrand breaks occur at a high rate (Marguet and Forterre, 1994; Grogan, 1998). The corresponding lesions could lead to lethal double-strand breaks, if not prevented or repaired in vivo. Therefore, it was suggested that hyperthermophiles must have evolved highly efficient mechanisms to protect and/or repair their DNA (Grogan, 1998). In support of this hypothesis, when Pyrococcus furiosus cells are exposed to 100∞C, their DNA is about 20 times more resistant to breakage than DNA from Escherichia. coli at the same temperature (Peak et al., 1995). Furthermore, passive protection of DNA might be provided by similar mechanisms as used to increase the Tm of the DNA double helix, i.e., high salt concentrations, and binding to proteins (see above). Indeed, it has been shown that the presence of Mg2+ and K+ protect double-stranded DNA from depurination, probably by directly stabilizing the N-glycosidic bond between the deoxyribose and the base (Marguet and Forterre, 1998). With respect to the formation of nucleoprotein complexes, archaeal histones are known to protect plasmid DNA against radiation (Isabelle et al., 1993). In spite of these well-established protection mechanisms, DNA in hyperthermophiles will almost certainly be damaged to a larger extent than DNA in mesophiles. A model organism for comparative research in this context is the radiation-resistant bacterium Deinococcus radiodurans. Both, g-irradiation and heat have been shown to induce double-strand breakage of DNA, which can be repaired efficiently by D. radiodurans. This capacity derives from multiple copies of its chromosome providing intact copies for repair by a DNA recombinase (Minton and Daly, 1995). In analogy, the chromosome of the archaeon Pyrococcus furiosus, after irradiationinduced fragmentation, was reassembled by the cells upon incubation at 95∞C (di Ruggiero et al., 1997). Open reading frames encoding homologues of RecA proteins involved in recombination repair in bacteria and eukarya were found in the archaeal genomes sequenced so far. Strong experimental evidence suggests that at least one of these homologs, FEN-1 from P. furiosus, is involved in double-strand breakage repair

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(di Ruggiero et al., 1999). Other than doublestrand breakage-repair, activities have been demonstrated in vitro for several other archaea. For example, Methanobacterium thermoautotrophicum is able to remove ultraviolet light-induced photoproducts, supposedly with a photolyase as catalyst (Kiener et al., 1989; Ögrünc et al., 1998). Furthermore, a T/G-selective DNA thymine Nglycosylase takes care of the mutagenic effect of hydrolytic 5-methylcytosine deamination (Horst and Fritz, 1996), while uracil-DNA glycosylases seem to be involved in the repair of cytosine deamination (Koulis et al., 1996); in addition, O6alkylguanine-DNA transferase activities were also found in hyperthermophiles (Skorvaga et al., 1998). On the other hand, MutL and MutS, which are used in all bacterial and eukaryal mismatchrepair systems, have not been found in any of the archaeal genomes so far. In summary, the present knowledge of the specific mechanisms by which hyperthermophilic microorganisms preserve the integrity of their genetic material is still incomplete. More information is needed about the intracellular salt concentrations and the DNA-binding and DNA-protecting proteins, to establish in vitro test systems that come as close as possible to the in vivo situation. Moreover, homologs of known bacterial and eukaryal repair enzymes from hyperthermophiles need to be characterized to identify their catalytic properties under physiological conditions. The ongoing genomesequencing projects will help identify the most promising candidates for this approach.

Adaptive Stabilization Mechanisms of Lipids and Membranes Living cells have a cytoplasmic membrane serving as a barrier between the cytoplasm and the environment. It consists of lipid layers with embedded proteins that generate specific and vital solute concentration gradients across the membrane. Penetration of small solutes through the lipid component of the membrane is caused either by active transport or passive diffusion. Being directly proportional to the thermal energy (kT), passive diffusion is accelerated with increasing temperature (Einstein, 1905; Einstein, 1906; van de Vossenberg et al., 1998). In hyperthermophiles, extreme temperature may lead to the breakdown of solute gradients. Therefore, their membranes need to be extremely thermostable, but they also require specific adaptive mechanisms to limit the permeability of ions. This holds especially for protons because of the essential role of proton gradients in energyrequiring processes such as ATP synthesis, active transport of specific solutes across the mem-

CHAPTER 1.7

brane, flagellar rotation, and maintenance of the intracellular pH and turgor (Albers et al., 2000).

Chemical Composition of Membrane Lipids At physiological temperatures, membrane lipids are in a liquid-crystalline state (Melchior, 1982), forming a suitable matrix for the attachment or integration of membrane proteins. The overall structure of the lipid membrane is conserved between eukarya, bacteria and archaea. The inner and outer hydrophilic surfaces, which are composed of polar headgroups, enclose the hydrophobic interior consisting of long hydrophobic hydrocarbon chains. At this point, the chemical composition of archaeal membranes has been found to be significantly different from the chemical composition of bacterial and eukaryal membranes. Both bacterial and eukaryal lipids have esters between glycerol and fatty acid chains (glycerol fatty acyl diesters), whereas the lipids of archaeal membranes are formed by ethers between glycerol (or another alcohol such as nonitol) and branched C20-hydrocarbon side chains (Langworthy and Pond, 1986). The side chains consist of repeated saturated isoprenoid units containing a methyl side group at every fourth carbon atom in the backbone. These methyl side groups restrict the mobility of the chains, thereby stabilizing them and restricting ion permeability (see below). The two hydrocarbon chains can be ether-linked to either one glycerol unit (forming a C20, C20-isopranyl glycerol diether = diphytanylglycerol diether = archaeol), or two glycerol units (forming a dibiphytanylglycerol tetraether = caldarchaeol; Fig. 8A and B). The archaeols are found in all archaea, whereas the caldarchaeols (and nonitol-caldarchaeols) are only found in thermophilic archaea. The caldarchaeols can be further modified by cyclopentane rings in the biphytanyl side chains (Fig. 8C–E). The caldarchaeols of thermophilic archaea are typically glycosylated at C3 and C6 of the glycerol and nonitol backbones, respectively. Probably, hydrogen bonds between the glycosyl headgroups stabilize the membrane structure by reducing lateral lipid mobility (van de Vossenberg et al., 1998; Daniel and Cowan, 2000). An unsaturated diether lipid was found in the archaeon Methanopyrus kandleri (Hafenbradl et al., 1993). This lipid, 2,3-di-O-geranylgeranylsn-glycerol, resembles terpenoids, but the consequences for membrane function are still unknown. Another type of unsaturated lipid was discovered in the psychrophilic archaeon Methanococcoides burtonii (Nichols and Franzmann, 1992). This lipid contains a double bond that can distort the short-range order of the membrane, thus allowing the necessary fluidity of the mem-

CHAPTER 1.7 Fig. 8. Archaeal lipid architecture. (A) Diphytanyl glycerol diethers, (B) dibiphytanyl diglycerol tetraethers, and (C–E) internal cyclization in dibiphytanyl diglycerol tetraethers. From Daniel and Cowan (2000).

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R =H (Caldarchaeol)

A R= O O

HO OH OH OH OH OH (Nonitolcaldarchaeol)

HO R OH

B O

O

O

O HO

R

C

OH

O O O O

HO

D

R

O

OH O

O HO

O

E O R OH

O HO

O O

brane to be adapted to the physiological low temperature (Suutari and Laakso, 1992).

Topology, Stability and Permeability of Membranes The glycerol-diester lipids of bacteria and eukarya form bilayer membranes. The same holds for the archaeol lipids of halobacteria and most other archaea growing under moderate conditions (Kates et al., 1993; Upasani et al., 1994; Kates, 1995). In contrast, the caldarchaeol lipids of the thermophilic and acidophilic archaea form monolayers spanning the entire membrane (de Rosa et al., 1991; Relini et al., 1996). In monolayers, two glycerol units are covalently linked by the phytanyl side chains, whereas in bilayers the glycerol units are noncovalently linked by hydrophobic interactions between the fatty acid side chains. As a consequence, monolayers have a diameter between 2.5 and 3.0 nm (Gliozzi et al., 1983), somewhat thinner than typical C18 glycerol-diester bilayers, but much more stable: Vesicles generated from Thermoplasma acidophilum ether lipids are more resistant to high temperature and surfaceactive agents than vesicles of bacterial dipalmitoyl phosphatidyl-choline (Ring et al., 1986).

Moreover, liposomes prepared from tetraether lipids from a number of archaea were shown to be extremely stable toward high temperature, alkaline pH and enzymatic degradation by phospholipases (Chang, 1994; Choquet et al., 1994). As has been mentioned, to guarantee energy production, membranes of all microorganisms, no matter whether they are psychro-, meso-, thermo- or hyperthermophilic, must provide an efficient barrier against the flux of protons. Liposomes prepared from lipids derived from a variety of organisms with different growth temperatures were compared for their proton permeabilities (van de Vossenberg et al., 1995). This study showed that, at the respective growth temperature, proton permeability was closely similar for the various liposomes (Fig. 9). This “homeoproton permeability adaptation” is reminiscent of the “corresponding states” observed for homologous pairs of enzymes from mesophiles and thermophiles, most of which were shown to exhibit comparable stabilities, flexibilities and activities at their respective physiological temperatures (Jaenicke, 1991b; Somero, 1995; Jaenicke and Böhm, 1998). As a logical consequence, at a given fixed temperature, the proton permeability of membranes is decreased with increasing temperature of adaptation, fol-

Proton permeability, k (S–1)

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R. Jaenicke and R. Sterner B

& ii B

10–1

CHAPTER 1.7

B

O C O

C O C

5x

i & &

0

B

B i

20

i

i

B

B

i

C

i i

40

O

C

O C

10–2

O

C

O C

i

60

80

100

Temperature (°C)

Fig. 9. The proton permeability of archaea and bacteria that live at different temperatures. At the respective growth temperatures, the proton permeability falls within a narrow range (gray bar). Thermotoga maritima and Bacillus stearothermophilus have higher permeabilities than those of other organisms. Both organisms overcome this problem differently. From Albers et al. (2000).

lowing the order: psychrophiles > mesophiles > thermophiles > hyperthermophiles. Various archaeol and caldarchaeol lipids were 6–120-fold less permeable to water, solutes, protons and ammonia than bacterial diphytanyl-phosphatidylcholine liposomes (Mathai et al., 2001). It was shown that the crucial factor ensuring low permeability are cyclopentane rings in the phytanyl side chains, which limit the mobility in the midplane hydrocarbon region. The substutitution of ether- for ester-bonds provides an additional barrier that specifically impairs the flux of protons. Bacterial thermophiles have membrane lipids rich in saturated fatty acids, which make the membranes more rigid and stable at high temperatures because stronger hydrophobic interactions are formed between saturated fatty acids compared with unsaturated ones (Brock, 2000). Other differences between membranes from mesophilic and thermophilic bacteria include alterations in acyl chain length, branching, and/ or cyclization (Tolner et al., 1998). Interestingly, the extremely thermophilic Thermodesulfobacterium contains lipids combining bacterial and archaeal properties; here, glycerol is ether-linked to a unique C17 hydrocarbon side chain along with some fatty acids instead of phytanyl side chains (Brock, 2000).

Adaptation of Membrane Structure and Function to Temperature Fluctuations Bacteria and archaea can grow over a wide range of temperatures. When facing environmental temperature shifts, most of them adapt the structure of their membranes to ensure constant stability and permeability. In archaea, as well as in

O

Fig. 10. Cyclization of the phytanyl chains of the S. solfataricus tetraether lipids. Only one of the phytanyl side chains is shown. The degree of cyclization increases from top to bottom. From Albers et al. (2000).

mesophilic and psychrophilic bacteria, this adaptation is achieved by adjusting the chemical composition of the lipids. Archaea adapt to low temperatures by decreasing the degree of saturation of their hydrocarbon side chains (Nichols and Franzmann, 1992), whereas they respond to high temperature by the cyclization of the side chains and by replacing diether to with tetraether lipids (de Rosa and Giambacorta, 1988; de Rosa et al., 1991; Yamauchi and Kinoshita, 1995): For Sulfolobus solfataricus and Thermoplasma, it was shown that the number of cyclopentane rings incorporated into the lipid diphytanyl side chains increase with growth temperature, this way rigidifying the membrane and limiting passive diffusion of small molecules (Mathai et al., 2001; Fig. 10). In Methanococcus jannaschii, a different mechanism is observed: here, increasing temperatures induce the change from diether lipids to the more thermostable tetraether lipids (Sprott et al., 1991). To investigate adaptive changes of membranes from bacteria, Bacillus subtilis was grown at the boundaries of its growth temperature (van de Vossenberg et al., 1999). The average lengths of lipid acyl side chains, the degree of saturation, and the ratio of iso- and anteiso-branched fatty acids increased with temperature. In accordance with the concept of homeoproton permeability adaptation, these modifications kept the proton permeability of the cytoplasmic membrane at a rather constant level. Likewise, in psychrophiles, the proton permeability is maintained at a constant level when the growth temperature is varied (van de Vossenberg et al., 1995). In contrast, in thermophilic bacteria such as Bacillus stearothermophilus and Thermotoga maritima, homeoproton permeability cannot be maintained, as their membranes become porous at high temperatures. Some moderately thermophilic bacteria can compensate for the high proton leakage by drastically increasing the respiration rate, and together with that, the rate of proton pumping (de Vrij et al.,

CHAPTER 1.7

1988). A different strategy is found in the moderate thermophile Caloramator fervidus, which, instead of the proton, uses the less permeable sodium ion as the main coupling component for energy transduction (Speelmans et al., 1993a; Speelmans et al., 1993b). In summary, a number of different mechanisms have been identified that keep membranes stable and functional at high temperatures. Archaea contain lipids with ether linkages between various alcohols and hydrocarbon side chains, in which cyclopentane rings are incorporated in a growth-temperature dependent manner. Thermophilic bacteria, which contain less stable ester lipids prone to proton leakage, evolved alternative strategies to maintain vital chemiosmotic gradients under physiological conditions. As the number of novel lipid structures constantly grows, more variations on these themes are to be expected.

Adaptive Stabilization Mechanisms of Proteins To fulfil their diverse functions, proteins from hyperthermophiles need to be in their native, folded state at temperatures around 100∞C. In contrast, most proteins from mesophiles are unfolded at ~50∞C (Fig. 2), often followed by irreversible aggregation and/or chemical damage (Jaenicke and Seckler, 1997). What are the structural determinants that render proteins from hyperthermophiles much more thermostable than their homologs from mesophiles? As mentioned in the section on “Stability of Biomolecules,” few additional favorable electrostatic or hydrophobic interactions suffice to shift DGstab of a protein from the mesophilic to the thermophilic temperature regime (Jaenicke and Böhm, 2001; Fig. 3B). In addition, proteins from hyperthermophiles are not only stabilized intrinsically, but also by extrinsic factors such as compatible solutes or molecular chaperones. What follows briefly summarizes our current knowledge of the intrinsic and extrinsic stabilization of hyperthermophilic proteins. For further details see (Jaenicke and Böhm, 2001; Petsko, 2001; Sterner and Liebl, 2001; Vieille and Zeikus, 2001).

Intrinsic Stabilization: There Are No General Rules In the section on “Stability of Biomolecules,” the electrostatic and hydrophobic interactions that stabilize proteins were discussed. Moreover, the contributions of enthalpy and entropy to the free energy gain caused by these interactions was pointed out. Pairwise comparisons of amino acid sequences and X-ray structures of homologous

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proteins from mesophiles, thermophiles and hyperthermophiles showed that one or more of these stabilizing interactions were more frequent or more pronounced in the thermophilic and hyperthermophilic variants. These additional stabilizing interactions can in principle occur at all levels, from primary to the quaternary structure (Jaenicke and Böhm, 1998; Daniel and Cowan, 2000; Vieille and Zeikus, 2001; Sterner and Liebl, 2001; Yano and Poulos, 2003). A large number of mutational studies have been performed to identify stabilizing interactions, which were frequently detected in hyperthermophilic proteins. To this end, selected amino acid residues were substituted by sitedirected mutagenesis, and the resulting changes in stability were measured. Instructive examples are the enzymes phosphoribosylanthranilate isomerase (PRAI) and indoleglycerol phosphate synthase (IGPS), which catalyze two successive reactions within tryptophan biosynthesis and adopt the frequently encountered (ba)8-barrel fold (Höcker et al., 2001; Wierenga, 2001). PRAI is monomeric in most mesophiles but dimeric in Thermotoga maritima (Sterner et al., 1996). The two identical monomers of Thermotoga maritima PRAI are associated via intimate hydrophobic contacts at the N-terminal faces of their central b-barrels (Hennig et al., 1997). By replacing a Phe residue at the monomer-monomer interface of T. maritima PRAI by a Glu residue, the hydrophobic interactions are weakened. As a consequence, the enzyme becomes monomeric and thermolabile, without losing its catalytic activity (Thoma et al., 2000; Fig. 11). The importance of increased association states for increased thermostability was also shown for ornithine carbamoyltransferase, which consists of four trimers in Pyrococcus furiosus, but only one in mesophiles. Gradual dissociation of dodecameric ornithine carbamoyltransferase from Pyrococcus furiosus into trimers, as induced by site-directed mutagenesis at subunit interfaces, led to a gradual decrease in thermal stability (Clantin et al., 2001). Indoleglycerol phosphate synthase is monomeric both in mesophiles and hyperthermophiles. However, IGPS from Sulfolobus solfataricus and T. maritima contain twice the number of potentially stabilizing ion pairs compared with E. coli (Hennig et al., 1995; Merz et al., 1999). Two T. maritima IGPS variants, which had one of these ion pairs disrupted by site-directed mutagenesis, showed significantly decreased thermostabilities (Merz et al., 1999). The stabilizing role of ion pairs was also proven by site-directed mutagenesis experiments performed with glyceralaldehyde-3phosphate dehydrogenase (GAPDH) from T. maritima, glutamate dehydrogenases from both Thermococcus litoralis and P. furiosus,

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α2

α2

Shortening of the loop between α2 and β2

and exchange Phe55Glu

α2

Fig. 11. Monomerization of the native homodimer of PRAI from Thermotoga maritima by rational design. Monomers were generated by shortening of the loops connecting helices a2 with strands b3 (in green), and by replacing the two Phe55 residues located close to the twofold symmetry axis (shown as a black dot) with glutamates (shown in stick format). The bound phosphate ions (red tetrahedrons) identify the active sites. The monomeric variants are catalytically as active as the dimer, but far more thermolabile. From Höcker et al. (2001), with permission.

3-isopropyl-malate dehydrogenase from Thermus thermophilus, rubredoxin from P. furiosus, and the archaeal histone from Methanothermus fervidus, and citrate synthase from psychrophiles to hyperthermophiles. The latter represents a good example for the whole spectrum of adaptive changes, including intersubunit ionic networks and varying states of association (Wrba et al., 1990; Tomschy et al., 1994; Pappenberger et al., 1997; Vetriani et al., 1998; Li et al., 2000; Nemeth et al., 2000; Strop and Mayo, 2000; Nordberg Karlsson et al., 2002, 2003; Bell et al., 2002, and references therein). The increased DGstab,70∞C

of the hyperthermophilic cold shock protein from T. maritima compared to its mesophilic counterpart from B. subtilis was shown to be largely due to Arg3, whose positive charge improves the global electrostatic potential of the protein (Perl and Schmid, 2001; cf. the section on “Stability of Biomolecules”). This result suggests that the optimum placement of charged groups on the surface of a protein is crucial for its thermostability (Xiao and Honig, 1999), a hypothesis that is strengthened by a number of other experimental studies (Grimsley et al., 1999; Loladze et al., 1999; Spector et al., 2000). In spite of these examples, in many cases the predicted stabilizing interactions (as deduced from pairwise mesophile-thermophile comparisons of sequences and structures) could not be verified experimentally. Therefore, it is still not possible to deduce general mechanisms that would lead to high protein thermostability. The reason for this shortcoming is the large number of neutral changes of amino acid residues and 3D structures that have accumulated during evolution without affecting protein stability (Böhm and Jaenicke, 1994; Arnold et al., 2001b). Based on this argument, large-scale structural comparisons of amino acid sequences and 3D structures, which reduce the large “phylogenetic noise,” are likely to provide more significant results. Such systematic comparisons are now possible owing to the growing number of complete genome sequences from mesophiles and hyperthermophiles, and the fast rate with which new X-ray structures become available. The amino acid compositions of a number of mesophiles and thermophiles were deduced from their genome sequences and compared in several systematic studies (Table 4). These comparisons allow the following conclusions. Hyperthermophilic proteins 1) contain a decreased content of uncharged polar amino acids, this way avoiding deamidation of Gln and Asn catalyzed by Thr and Ser (Wright, 1991; Haney et al., 1999; cf. the section on “Biochemical Limitations at High Temperature”), 2) show an increased content of the charged amino acids Glu and Asp, a significant fraction of which may be involved in stabilizing ion pairs at the protein surface (see above; Haney et al., 1999; Cambillau and Claverie, 2000), and 3) are on avarage significantly smaller than their mesophilic homologs (Chakravarty and Varadarajan, 2000), presumably owing to shorter solvent-exposed surface loops (Thompson and Eisenberg, 1999) or extensions at the N- and/or C-terminal ends (Fig. 12). Upon unfolding, small proteins show a smaller heat capacity change (DCp) than large proteins (Murphy and Freire, 1992; Myers et al., 1995); a decrease in DCp flattens the DGstab versus T profile and leads to an increase in Tm (Fig. 3B).

CHAPTER 1.7

Life at High Temperatures

The three-dimensional structures of proteins from mesophiles and thermophiles were compared in a number of comprehensive studies. From a non-redundant dataset of high-quality Xray structures of protein subunits from mesophiles, thermophiles and hyperthermophiles, it revealed that the increase in intrinsic stability was paralleled by more ion pairs (apart from slight differences with respect to cavities), hydrogen bonds, secondary structure content and polarity of surfaces (Szilagyi and Závodszky, 2000; Table 5). A similar study suggested that ion pairs and side chain-side chain hydrogen bonds are more frequent in thermophilic than in mesophilic proteins (Kumar et al., 2000a; 2000b). There was no evidence for significant differences with respect to compactness, hydrophobicity, polar and nonpolar surface area, protein size, and number of Pro residues in loops; however, thermophilic proteins appeared to have a higher fraction of residues in a-helices. Two further investigations confirmed that the a-helices of thermophilic proteins show increased stability, mainly due to the higher intrinsic helical propensities of the amino acids involved (Petukhov et al., 1997; Facchiano et al., 1998). Two systematic comparisons of lactate dehydrogenases (LDH) and triosephosphate isomerases (TIM) from psychrophiles, mesoTable 4. Change in amino acid composition going from proteins of mesophiles to proteins of thermophiles. Amino acid Ile Glu Arg Lys Pro Tyr Ala Trp Leu Cys Phe Asp Val His Gly Met Gln Thr Asn Ser

Gains

Losses

Ratio

Net change

Change, %

842 739 383 789 167 224 504 23 560 72 200 429 666 80 201 174 158 336 313 271

658 562 214 620 96 177 458 11 548 69 202 432 670 92 264 248 234 431 481 664

1.28 1.31 1.79 1.27 1.74 1.27 1.10 2.09 1.02 1.04 0.99 0.99 0.99 0.87 0.76 0.70 0.68 0.78 0.65 0.41

184 177 169 169 71 47 46 12 12 3 -2 -3 -4 -12 -63 -74 -76 -95 -168 -393

9.5 9.1 16.5 8.3 7.0 5.8 2.8 8.3 0.6 0.9 -0.3 -0.2 -0.2 -2.8 -3.4 -11.3 -13.1 -8.4 -15.9 -31.7

Abbreviations: Ile, isoleucine; Glu, glutamic acid; Arg, arginine; Lys, lysine; Pro, proline; Tyr, tyrosine; Ala, alanine; Trp, tryptophan; Leu, leucine; Cys, cysteine; Phe, phenylalanine; Asp, aspartic acid; Val, valine; His, histidine; Gly, glycine; Met, methionine; Gln, glutamine; Asn, asparagine; Ser, serine. Data from Haney et al. (1999).

193

philes and hyperthermophiles revealed positive correlations between thermostability and the number of intra-subunit (LDH) and intersubunit ion pairs (TIM), respectively (Auerbach et al., 1998; Maes et al., 1999). The results of the cited mutational studies, and those of the systematic and comprehensive comparisons between the amino acid sequences and 3D structures of psychrophilic, mesophilic and thermophilic proteins can be summarized as follows: Owing to the small differences between DGstab of hyperthermophilic and mesophilic proteins (Matthews, 1993; Matthews, 1996; Jaenicke and Böhm, 1998), attempts to find a unifying set of rules of stabilization must fail. The structural features that characterize some of the known hyperthermophilic proteins are increased numbers of hydrogen bonds, higher packing densities and a-helical contents, improved hydrophobic interactions, optimized surface areas, decreased volumes, fewer cavities, and a shortening of the polypeptide chains. Attempts to define the relative significance of these many different factors by counting their frequency in comprehensive comparative studies led to four major contributions: 1) stabilized a-helices, 2) decreased entropy of the unfolded state by increased numbers of Pro and b-branched amino acid residues, 3) decreased content of chemically labile polar amino acid residues, and in particular, 4) increase in the number of optimized ionic interactions (Sanchez-Ruiz and Makhatadze, 2001). The latter finding is in accordance with theoretical work suggesting ion pairs are more stabilizing at high than at low temperatures and might therefore be crucial for the stability of hyperthermophilic proteins (Elcock and McCammon, 1997; Elcock, 1998; De Bakker et al., 1999). It is important to note that the stabilizing effect of a given ion pair depends on its structural context. Ion pairs that connect N- and C-termini in IGPS and GAPDH from T. maritima contribute significantly to thermostability, probably by preventing the fraying of the N- and C-termini, which might initiate thermal denaturation (Pappenberger et al., 1997; Merz et al., 1999). Also, for entropic reasons, clusters of ion pairs are likely to be more stabilizing than individual ion pairs (Yip et al., 1995; Yip et al., 1998). Although our knowledge of the structural basis of high intrinsic protein thermostability is still incomplete, considerable operational progress has been achieved in the last years, especially in the first successful examples of rational or semi-empirical improvements of protein thermostability (Malakauskas and Mayo, 1998; van den Burg et al., 1998). An alternative approach to improve protein thermostability is “directed molecular evolution” (Wintrode and Arnold, 2000; Arnold et al., 2001b). It mimics the

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CHAPTER 1.7

saldh blldh

ssldh bsldh

Icldh

300 125 100 175 75 225 250 75 200 150 25 50

tmldh

Fig. 12. The three-dimensional structures of lactate dehydrogenases (ldh) from hyperthermophiles, on the one hand, and mesophiles as well as a cold-blooded fish, on the other, are practically isomorphous, with root-mean-square (r.m.s.) differences below 2.4 Å. Comparisons of the 2–3 Å resolution crystal structures of the various homologs with the enzyme from Thermotoga maritima (as the reference; in gray) show that the increase in thermostability is parallelled 1) by a reduction in the length of the C-terminal extension, 2) by an increase in compactness of the tetrameric assembly, and 3) by the presence of an additional “thermohelix” (aT) in each of the subunits of the hyperthermophile enzyme. The shift from blue to red in the figure characterizes the temperature range of the organisms from which the various enzymes were isolated The corresponding abbreviations and physiological Topt-values refer to: sa, Squalus acanthias (dogfish, ~10∞C); ss, Sus scrofa (pig, 37∞C); lc, Lactobacillus casei (~30∞C); bl, Bifidobacterium longum (~40∞C); bs, Bacillus stearothermophilus (~65∞C) and tm, Thermotoga maritima (~80∞C). For details, see Auerbach et al. (1998).

natural evolution process by applying iterative rounds of random mutagenesis and selection (or screening) of stabilized protein variants. Given an appropriate selection or screening system, this approach is generally applicable because it

does not require specific knowledge of the structure of the protein to be stabilized. Moreover, directed evolution is instructive because it is unbiased and may provide stabilizing amino acid exchanges at positions in the protein that were

Table 5. Systematic comparison of the structures of proteins from mesophiles, thermophiles and extreme thermophiles.

Property Cavities Hydrogen bonds Ion pairs Secondary structure Polarity of surfaces

Number Volume Area Number Unsatisfied 35

a

The temperatures given for a psychrophile are taken from Morita (1975). Taking into account the temperature variations for the growth of microorganisms in refrigerated food reported in the literature, the lower growth limit of psychrotolerants in Table 1 has been set to 7°C.

Table 2. Selection of psychrotolerant and psychrophilic species described between 2001 and beginning of 2005. Species

TT

Isolated from

Reference(s)

Algoriphagus antarcticus Alkalibacterium olivoapovliticus Alkalibacterium psychrotolerans Alteromonas stellipolaris Arthrobacter psychrophenolicus Bacillus psychrodurans Bacillus psychrotolerans Carnobacterium pleistocenium Chromohalobacter sarecensis Clostridium sp. PXYL1 Dietzia psychralcaliphila Flavobacterium frigidarium Flavobacterium frigoris Geopsychrobacter electrodiphilus Gillisia limnaea Glaciecola polaris Halomonas boliviensis Hyphomonas aff. jannaschiana Lactovum miscens Marinilactibacillus piezotolerans Marinobacter aff. aquaeolei Marinomonas ushuaiensis Methanogenium marinum Methanosarcina mazei Mycobacterium psychrotolerans Paenibacillus antarticus Pseudomonas alcaliphila Pseudomonas antartica Pseudomonas psychrophila Pseudomonas psychrotolerans Psychrobacter marincola Psychrobacter maritimus Psychrobacter nivimaris Psychrobacter proteolyticus Psychrobacter salsus Psychrobacter sumarinus Psychromonas profunda Rhodoferax ferrireductans Sejongia antartica

pp pt pt pt pp pt pt pt pt pp pp pp pp pt pp pt pp pp pt pt pp pp pt pt pt pt pp pp pp pt pp pt pt pt pt pp pp pt pt

microbial mats in Antartic lake Olive wash-waters fermented polygonum indigo Antartic sea water alpine cave Soil, Egypt Soil, Germany permafrost, Alaska saline Andean region Cattle manure digester, India Fish-processing plant, Japan Antarctica microbial mats in Antartic lake marine sediment fuel cell microbial mats in Antartic lake Artic ocean Bolivian hypersaline lake Deep sea acidic forest soil deep sub-seafloor sediment Deep sea coastel sea water, Argentina Marine sediment, Alaska Tundra pond water near uranium mine Antartic sediment Seawater, Hokkaido, Japan microbial mats from Antartica Food storage room, Japan vetinary hospital, Vienna Sea water coastal sea ice, sea of Japan Southern Ocean Antarctic krill fast ice, Adelie Land, Antartica Sea water Deep Atlantic sediment Marine sediment, United States terrestrial samples, Antartica

Van Trappen et al., 2004c Ntougias and Russell, 2001 Yumoto et al., 2004 Van Trappen et al., 2004a Margesin et al., 2004 Abd El-Rahman et al., 2002 Abd El-Rahman et al., 2002 Pikuta et al., 2005 Quillaguaman et al., 2004a Akila and Chandra, 2003 Yumoto et al., 2002 Humphry et al., 2001 Van Trappen et al., 2004d Holmes et al., 2004 Van Trappen et al., 2004e Van Trappen et al., 2004b Quillaguaman et al., 2004b Edwards et al., 2003 Matthies et al., 2004 Toffin et al., 2005 Edwards et al., 2003 Prabagaran et al., 2005 Chong et al., 2002 Simankova et al., 2003 Trujillo et al., 2004 Montes et al., 2004 Yumoto et al., 2001b Reddy et al., 2004 Yumoto et al., 2001a Hauser et al., 2004 Romanenko et al., 2002 Romanenko et al., 2004 Heuchert et al., 2004 Denner et al., 2001 Shivaji et al., 2004 Romanenko et al., 2002 Xu et al., 2003c Finneran et al., 2003 Yi et al., 2005

Abbreviations: TT, thermal type; pt, psychrotolerant; and pp, psychrophilic.

free floating cells revealed significant differences in the protein response (Perrot et al., 2001). What one would define as a specific cold shock stimulon is therefore dependent on the experimental procedures. To our knowledge, virtually no report deals with the cold shock response of

stationary phase cells, though many bacteria spend most of their lifetime in stationary phase (Kjelleberg, 1993). Although low temperature induces an adaptation of many cellular components, e.g., the membrane composition, the supercoiling of the DNA,

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and the transcriptome (see the section Cold Shock and the Degradation in this Chapter), the most severe problem seems to be initiation and translation of bulk mRNA at low temperatures (Broeze et al., 1978; Jones and Inouye, 1994). Shifting a culture of Escherichia coli from 37°C to 8°C or below resulted in polysomal run-off and accumulation of free ribosomes (Broeze et al., 1978; Xia et al., 2002). Other stresses, e.g., some antibiotics (VanBogelen and Neidhardt, 1990), dilution of a culture with fresh medium (Brandi et al., 1999a; Brandi et al., 1999b), an upshift in the concentration of nutrients (Yamanaka and Inouye, 2001a), diauxic lag (Novotna et al., 2003), oxidative stress (Smirnova et al., 2001b), hydrostatic pressure (Wemekamp-Kamphuis et al., 2002), or exposure to colicin E9 (Walker et al., 2004) can mimic a cold shock response or at least induce the cold shock stimulon to some extent (Wick and Egli, 2004). Therefore it could be hypothesized that every event stopping or stalling the ribosomes leads to an induction of the cold shock response (Walker et al., 2004). This is most obvious for the cold shock response itself (Gualerzi et al., 2003), the use of certain antibiotics affecting the translational speed (VanBogelen and Neidhardt, 1990), or after nutrient upshift (Brandi et al., 1999a) in which many new mRNAs are synthesized to adapt to the new nutrition provided. Chilling of bacterial cells affects their viability. In Bacillus subtilis it was shown that death after cold shock is not only due to a passive event (a reaction velocity decrease and outrun of energy), but also to translocation of a DNase (YokF) from the periplasm into the cytoplasm attacking the DNA of the cell. This partly resembles apoptosis in eukaryotic cells (Sakamoto et al., 2001). Most work on cold shock was done with the mesophiles E. coli and B. subtilis. This article, therefore, reports mainly on these two organisms. The reader may consult the detailed review by Weber and Marahiel (2003) about bacterial cold shock responses. Besides cold shock proteins, some cold adaptation proteins have been described. However, much less is known about the permanent response of bacteria towards low temperature. This work is summarized in the section Cold Adaptation.

Major Cold Shock Proteins: CspA–CspI In most free-living bacteria, a cold shock protein (Csp) family has been identified (Francis, 1997) and displays homology to CspA, first discovered in E. coli (Jones et al., 1987); reviewed in Ermolenko and Makhatadze (2002). Since CspA has the highest induction level, these proteins are often termed “major cold shock protein(s)”

CHAPTER 1.8

Fig. 1. Regulation of bacterial cold shock responses as a multiple filter model. The arrows indicate the flow of genetic information from DNA (via RNA polymerase, top) to protein (bottom), in turn affecting via a feedback circuit response (eventually conducted by effector molecules) this genetic flow. The T-arrows have to be read as “modulates activity of.” The filter systems are boxed in square boxes. These filters are integration systems reacting directly or indirectly to temperature changes. Adapted from Weber and Marahiel (2003).

(MCSPs; Goldstein et al., 1990; Etchegaray and Inouye, 1999b; Lopez and Makhatadze, 2000). However, note that many parasitic or pathogenic bacteria do not contain such a protein family (e.g., Chlamydia trachomatis, Helicobacter pylori, Mycoplasma sp. and others (Yamanaka, 1999a) and some psychrotrophic bacteria, as Aeromonas hydrophila, may not respond with a “typical” cold shock response (Imbert and Gancel, 2004). After a downshift from 37°C to 10°C, the major cold shock proteins of E. coli (CspA, B, G and I) are induced. The Csps reach 13% of the total protein synthesis, and synthesis of CspA is increased 30-fold under certain circumstances (Jones et al., 1987; Goldstein et al., 1990; Lee et al., 1994; Thieringer et al., 1998; Etchegaray and Inouye, 1999b; Wang et al., 1999; Gualerzi et al., 2003). In the psychrotolerant Yersinia enterocolitica, a CspA tandem has been discovered which may lead to a higher rate of CspA synthesis (Neuhaus et al., 1999). A first hypothesis formulated on the basis of CspA’s abundance after a cold shock was that this protein may function as an antifreeze protein.

CHAPTER 1.8

Life at Low Temperatures

213

RNAP

CSP

Cold shock Artificial decrease in CSP levels other stresses

CSP levels must increase to compensate for higher stability of secondary structures in RNA

Fig. 2. Model for the function of cold-shock proteins (CSPs) as RNA-chaperones that couple transcription to translation of mRNA. During growth at 37°C, CSPs bind to mRNA as it protrudes from the RNA-polymerase complex (RNAP) and maintain the RNA in a linear form. The ribosome then displaces CSPs, which have only low affinity for RNA, and initiates translation. Accordingly, an artificial decrease in the CSP concentration would lead to the formation of secondary structure in RNA and prevent translation. After cold shock or other stresses (e.g., carbon starvation), an increase in the CSP concentration is needed to counterbalance the increased stability of RNA secondary structure. Redrawn after Graumann and Marahiel (1998).

Although a cold adaptation period preceding freezing enhances freezing tolerance (Thammavongs et al., 1996); Kim and Dunn, 1997; Kim et al., 1998), the role of MCSPs in that protection has not been demonstrated unequivocally (Wouters et al., 1999). Apparently, CspA is-at least in part-an mRNA chaperone, opening the secondary structures of mRNAs at low temperature, an alteration which helps the ribosomes to function after a cold shock (Jiang et al., 1997); Fig. 2). Besides CspA, E. coli contains a family of highly similar Csps, containing eight other members: CspB consists of 71 amino acids (aa) and is 79% identical to CspA. Similarly, CspC contains 69 aa and has 70% identity to CspA; CspD (74 aa) has 45% identity; CspE (69 aa) has 70% identity; CspF (70 aa) has 44% identity; CspG (70 aa) has 73% identity; CspH (70 aa) has 47% identity; and CspI (70 aa) has 70% identity (Lee et al., 1994; Yamanaka et al., 1994; Yamanaka et al., 1998; Nakashima et al., 1996; Wang et al., 1999). Also, Bacillus cereus contains a family of MCSPs (Mayr et al., 1996). All these different CspA homologs are believed to be stress adaptation proteins for different tasks, but the coldinducible Csps can replace each other to some extent (Graumann et al., 1997; Yamanaka et al., 1998; Gualerzi et al., 2003). CspA is induced after a cold shock from 30°C down to 10°C, CspB and CspG occur between 20°C and 10°C, and CspI occurs below 15°C (Wang et al., 1999). A quadruple deletion mutant missing CspA, CspB, CspE and CspG was cold sensitive and formed filamentous cells at 15°C. This phenotype was suppressed by overexpression of each member of the cold-shock protein family except CspD, which causes lethality (Phadtare and Inouye, 2004b; Xia et al., 2001b). A different function of CspA and CspD was supported by another line of evidence. Green fluorescent protein (GFP)

fusions were found in the nucleoid in the case of CspD, and in a polar position of the cell in the case of CspA (Giangrossi et al., 2001a), however, in Pseudomonas Csp seems to be distributed evenly in the cytosol of the cell (Khan et al., 2003). Most cold inducible MCSPs have an unusually long mRNA leader region of 156–256 bp upstream of the translational start. An exception is the mRNA leader region of the cspH gene from Salmonella enterica, which is only 55 bp long (Kim et al., 2001). Another exception was reported recently, the cold-inducible CspAhomolog CspV from Vibrio cholerae, which exhibits a leader of only 12 bp (Datta and Bhadra, 2003). The CspA molecule is small (7.4 kDa and 70 aa), acidic (pI 5.92), and very hydrophilic (Goldstein et al., 1990). A remarkable feature is the high sequence similarity of the bacterial major cold shock proteins to eukaryotic Y-box factors, including human YB-1 (which is 44% identical with CspA) and frog FRG Y1/2 (Didier et al., 1988; Tafuri and Wolffe, 1990; Lee et al., 1994). These are domains of DNA or RNA-binding motifs which bind to a specific regulatory sequence called the “Y-box motif,” ATTGG/ CCAAT (Wolffe, 1994), and are therefore designated “cold-shock domains” (CSDs; Karlson and Imai, 2003). Interestingly, a protein with a similar fold, initiation factor 1, can complement in a B. subtilis cspB cspC double mutant (Weber et al., 2001a). Figure 3 shows the three-dimensional (3D) structure of CspB from B. subtilis (Schindelin et al., 1992; Schindelin et al., 1993; Schindelin et al., 1994; Schnuchel et al., 1993), which is similar to that CspA from E. coli (Schindelin et al., 1994; Feng et al., 1998). It consists of five antiparallel β-sheets, which form a barrel. The RNP-1 motif KGFGFI (Landsman, 1992; Lee et al., 1994) and RNP-2 motif VHVHF

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CHAPTER 1.8 C

N K7 β1

β5

β4

β2

β3 F 17 F 27 F 15

H 29

K 13

Fig. 3. Three-dimensional structure of CspB from B. subtilis features five antiparallel β-sheets, which form a barrel. From Graumann and Marahiel (1996a).

(Landsman, 1992; Schnuchel et al., 1993) contain most of the aromatic residues. They are exposed to the water phase and interact with singlestranded (ss) DNA (Schröder et al., 1993; Newkirk et al., 1994). If the aromatic residues in the RNP-1 or RNP-2 are replaced as a result of mutations, binding of ssDNA containing the Y-box motif is abolished (Schröder et al., 1995). CspB is very similar to CspA, the time frame of its induction follows closely that of CspA (Etchegaray et al., 1996). The promoter regions of cspA, cspB and cspG display similar features with similar promoter sequences and the same unusually long leader region of about 156–256 bases (Nakashima et al., 1996; Datta and Bhadra, 2003). The function of CspH needs to be elucidated (Yamanaka et al., 1998); however, in Salmonella enterica, cspH was described as cold inducible (Kim et al., 2001). Among the non-cold-inducible MCSPs, CspC is rather highly expressed at normal growth temperature (37°C) and its level remains unchanged after a cold shock (Lee et al., 1994). Also, CspC was found to be a multicopy repressor of the mukB106 mutant (Yamanaka et al., 1994). This mutant has a defect in chromosomal partitioning (Niki et al., 1991). CspD is induced at the onset of the stationary phase and inversely dependent on growth rate or glucose starvation. Guanosine3′-diphosphate-5′-(tri)diphosphate, collectively abbreviated (p)ppGpp, is a positive regulator for CspD (Yamanaka and Inouye, 1997; Yamanaka et al., 1998). Because overproduction of CspD is lethal, the presence of overproducing cells is

indicated by a typical morphology, which is due to impaired DNA replication. CspD inhibits effectively both the initiation and the elongation of minichromosome replication in vitro (Yamanaka et al., 2001c), and it is switched off after cold shock (Lee et al., 1994). CspD has been found associated with the nucleoid in E. coli (Giangrossi et al., 2001a). The function of CspF is unknown and needs to be elucidated (Yamanaka et al., 1998). CspE is abundantly produced at 37°C, but a cspA deletion mutant also has higher levels of CspE in the cold. Originally, CspE was found as a multicopy repressor of the mukB106 mutant gene that codes for a protein, which, together with CspC, plays a role in chromosomal partitioning (Yamanaka et al., 1994). Later, CspE was found to interact with nascent RNA in transcription complexes, causing antitermination. The latter function is coupled to the nucleic acid melting abilities of this protein (Phadtare et al., 2002a; Phadtare et al., 2002b; Phadtare et al., 2004c). Furthermore, it binds to the Y-box motif and functions as a repressor for cspA at 37°C through an interaction with the transcription elongation complex (Bae et al., 1999). Recently, it was discovered that CspE binds to poly(A) tails of mRNAs (which is a decay signal) and subsequently impedes the 3′ to 5′ exonucleolytic decay by polynucleotide phosphorylase (PNPase). CspE also inhibits both internal cleavage and poly(A) tail removal by RNase E, thus stabilizing mRNA (Feng et al., 2001). CspE was also found to be important in radiation resistance of E. coli (Chattopadhyay, 2002; Mangoli et al., 2001). All evidences taken together imply that CspE is a regulator also important for translational fidelity of DNA in cold environments, and for DNA condensation and partitioning during growth (Mangoli et al., 2001; Sand et al., 2003). The most prominent Gram-positive bacterium examined with respect to cold shock is the mesophile B. subtilis (Weber and Marahiel, 2002) extensively reviewed the cold shock response of this organism. Briefly, after a cold shock from 37°C to 15°C, protein synthesis resumed 2 h later. During the adaptation, CspB (which is a homolog to CspA of E. coli) is induced and remains at higher than pre-cold shock levels (Willimsky et al., 1992; Kunklova, 1995; Graumann et al., 1996b). A cspB::lacZ fusion showed a sevenfold induction after cold shock from 37°C to 10°C. In addition to CspB, CspC and CspD of B. subtilis are homologs; CspC is also cold inducible (Graumann et al., 1997) but differs slightly from CspB since the CspC increases more rapidly. Interesting genome-wide transcriptional profilings of the B. subtilis cold shock response were conducted by Kaan et al. (2002) and Beckering et al. (2002), the former

CHAPTER 1.8

Life at Low Temperatures

study describing genes not only induced but also repressed after cold shock in this organism. In B. subtilis, CspB and CspC not only participate in the cold shock response, but also act as major stationary-phase induced proteins. This illustrates the broad functionality of these Csps in cellular physiology (Graumann and Marahiel, 1999a). In Anabaena sp., no MCSP could be detected. However, an RNA helicase CrhC was found to be induced after cold shock (Chamot et al., 1999; Chamot and Owttrim, 2000). This helicase is completely membrane bound and mainly polar localized in this organism (El-Fahmawi and Owttrim, 2003). Interestingly, E. coli CspA has also been found in a polar position, but it remains in the cytoplasm (Giangrossi et al., 2001a). Even though the function of CrhC is not completely clear, the RNA unfolding abilities of both proteins, CrhC and CspA, seems to be needed in polar positions. In Synechococcus sp., a heat shock protein (Hsp)90 homolog, HtpG, was found to be heat and cold shock inducible (Hossain and Nakamoto, 2003).

Regulation of the Major Cold Shock Proteins The regulation of CspA induction in E. coli after cold shock is rather complex and not yet fully understood. Transcriptional and posttranscrip-

Sensors

Low positive temperature (from 4 to 16°C)

tional regulation of cold shock genes, including the MCSPs, was reviewed in detail by Gualerzi et al. (2003), and an overview of some aspects is given in Fig. 4. The cspA gene exhibits an unusually long leader sequence. The major transcription start +1 is located 159 bp upstream from the translational starting point. The promoter seems to be σ-70 dependent, since the –35 region (TTGCAT) and the –10 region (CTTAAT) are found to be similar to a σ-70 consensus sequence (TTGACA for the –35 and TATAAT for the –10 (Qoronfleh et al., 1992; Tanabe et al., 1992). Other regulatory elements in the gene sequence of cspA include the cold box (Fang et al., 1998; Jiang et al., 1996b), the upstream (UP) element, the downstream box (Mitta et al., 1997), the upstream box (Yamanaka et al., 1999c), and others (Yamanaka, 1999a). An overview of the features of the cspA gene sequence is given in Fig. 5. The 5′ end of the cspA mRNA contains a regulatory sequence (cold box), which stabilizes the mRNA at low temperature, enabling cold shock induction (Xia et al., 2002). The consensus cold box sequence (5′ UGACGUACAGA) is found in cspA, cspB and csdA (Jiang et al., 1996b). However, if the 5′ end of cspA containing this cold box is overproduced, the expression of cold shock genes is no longer transient, and the synthesis of bulk proteins is impaired (Jiang et al., 1996b; Xia et al., 2002). Also, the cessation of regrowth after cold shock is prolonged. This fits

Molecular regulation of the response

Signal

Translational control

Ribosome (p)ppGpp (?)

215

CspA Ribosome

CsdA RbfA IF2 PNP

5’ - CCAAT-Cold Shock Genes - 3’ Cold shock DNA Transcriptional control

H-NS GyrA

Cytoplasmic membrane RecA NusA

Other CSPs synthesis

DNA

Concomitance or Causality Negative temperature (from –16°C to –80°C)

Fig. 4. Some aspects of the cold shock response. From Panoff et al. (1998).

Cryotolerance

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CHAPTER 1.8 +1 (major) Ybox-motif

–35

–10

+1 (minor)

UP-element

cgattaatcataaatatgaaaaataattgttgcatcacccgCCAATgcgtggcttaatgcacatcaAcggtttga primary promoter +1 (secondary)

cold box

cgtacagaccattaaagcagtgtagtaaggcaagtcccttcaagagttatcgttgatacccctcgtagtgcacat induction enhancing mutations

secondary promoter proposed RNaseE cut site

tcctttaacgcttcaaaatctgtaaagcacgccatatcgccgaaaggcacacttaattattaAAGGtaatacact upstream box RNP-1

b1

+160

Shine-Dalgarno-S.

b2

M S G K M T G I V K W F N A D K G F G F I T P D D atgtccggtaaaatgactggtatcgtaaaatggttcaacgctgacaaaggcttcggcttcatcactcctgacgat start codon

downstream box b3

RNP-2

G S K D V F V H F S A I Q N D G Y K S L D E G Q K ggctctaaagatgtgttcgtacacttctctgctatccagaacgatggttacaaatctctggacgaaggtcagaaa b5

b4

gtgtccttcaccatcgaaagcggcgctaaaggcccggcagctggtaacgtaaccagcctgtaatctctgcttaaa stop codon terminator

agcacagaatctaagatccctgccatttggcggggatttttttatttgttttcaggaaataaataatcgatcgcg inverted repeats Fig. 5. Overview of the features of the cspA gene and the CspA protein.

nicely with the observation that cspA mRNA in excess is poisonous to the cell. The robust translatability of cspA mRNA depends on initiation, and the ribosome appears to be preadapted to translate cspA mRNA (Etchegaray and Inouye, 1999b; Xia et al., 2001a). Since overproduction of CspA together with the overproduction of the 5′-end restores the normal cold shock response, CspA itself probably interacts with the cold box (Jiang et al., 1996a). Furthermore, Giuliodori et al. (2004) could repeat the preferential translation of cold-shock mRNAs after cold shock in vitro. Apparently, cold-shock 70S ribosomes display some translational selectivity for MCSP mRNAs. The trans-acting factors involved are 1) CspA itself (increasing translatability of mRNA in the cold) and 2) the cold shock-induced stoichiometric imbalance between the initiation factors IF1, IF2 and IF3, on the one hand, and the ribosomes, on the other. Possible cis-acting elements discussed are the secondary or tertiary structures of the unusual long 5′ leader sequences of MCSP mRNAs. Furthermore, in addition to CspA-mediated autoregulation, a repressor for cspA was found, which turned out to be CspE. CspE is abundantly produced at 37°C, and in a cspE mutant, cspA is derepressed (Fang et al., 1998). In vitro,

CspE and CspA cause transcriptional pausing just behind the cold box of cspA, and CspA production is inhibited by addition of CspE to the translating ribosomes (Bae et al., 1999; Phadtare and Inouye, 1999). Mutational analysis of the 5′-untranslated leader of cspA showed another element to be involved in regulation of CspA. A deletion of a few bases upstream from the Shine-Dalgarno sequence (SDS) decreased the CspA amount more than 10-fold. It turned out that a 13-bp sequence located 11 bp upstream of the SDS is conserved in the cold-inducible genes cspA, cspB, cspG and cspI. This element was designated the “upstream box,” and it is speculated that this region may form different secondary structures at different temperatures, leading to an efficient translation at low temperatures or nearly zero translation at higher temperatures (Yamanaka et al., 1999c). Another element in CspA induction is the socalled “downstream box” (DB; Mitta et al., 1997). This element is found downstream of the ATG start codon of some cold shock genes and, according to its proponents, should be able to anneal to a complementary anti-downstream box at the 16S rRNA, thereby enhancing translation initiation. The existence of the downstream box

CHAPTER 1.8

has been disputed. Sprengart et al. (1996), Etchegaray and Inouye (1999a), Etchegaray and Inouye (1999c), Etchegaray and Inouye (1999d), Mironova et al. (1999), and Xia et al. (2001a) are in favor, while O’Connor et al. (1999), Resch et al. (1996), Bläsi et al. (1999), La Teana et al. (2000), and Rocha et al. (2000) are against. Apparently, Moll et al. (2001) finally rejected the concept of the downstream box.

Cold Shock and the Degradosome Recently, the mRNA-decay machinery of bacteria came into focus. In general, adaptation to low temperature after cold shock includes the establishment of a new equilibrium of the transcriptome following changes in transcription and mRNA decay rates, both of which are important for gene regulation in bacteria. The mRNA content of the cell is therefore not only regulated by cold shock induction or repression of certain genes, but also by stabilization or destabilization (depending on the specific mRNA and on the usage of different subsets of RNases; Mohanty and Kushner, 2003; Polissi et al., 2003). The subsets include PNPase and RNase H, which are cold shock induced, and RNase II or RNase E, which are not (Cairrão et al., 2003). The induction of CspA is mainly due to an increase in mRNA stability. Its half-life is 12 s at 37°C, but between 15 min and 30 min at 15°C in E. coli (Tanabe et al., 1992; Jiang et al., 1993; Fang et al., 1997; Gualerzi et al., 2003). If the coding region of cspA is fused to the constitutive promoter lpp, it is still cold inducible. This observation is explained by a strong vulnerability of the transcript to RNase E degradation at 37°C. Even if the cspA promoter is turned on constitutively, CspA can only be synthesized if the transcript is stabilized, perhaps by CspE (Fang et al., 1997; Feng et al., 2001). As in E. coli, the transcripts of cspB and cspC in B. subtilis are also dramatically stabilized, having a half-life of 1 min at 37°C and more than 30 min at 15°C (Kaan et al., 1999). A similar observation was made in Rhodobacter capsulatus with a cspA transcript half-life of around 4 min at 32°C and 47 min at 10°C (Jäger et al., 2004). Downregulation of MCSP mRNA is an important step, at least in enterobacteria, before growth can resume. This phenomenon is mainly due to the exceptionally strong ability of MCSP mRNAs to initiate at the ribosome. Therefore MCSP mRNA outcompetes bulk mRNA and thus prevents growth (Neuhaus et al., 2000b; Xia et al., 2001a; Yamanaka and Inouye, 2001b). In the above-mentioned PNPase-deficient strains, the decay of cspA mRNA is delayed, subsequently preventing re-growth (Neuhaus et al., 2000b). In Yersinia enterocolitica, the cspA tan-

Life at Low Temperatures

217

dem mRNA is cleaved at multiple specific cut sites, with an AGUAAA consensus (termed “cold shock cut box”) to downregulate the MCSP mRNA. After these initial cleaving steps, the fragments are removed rapidly and growth can resume (Neuhaus et al., 2003). Cleavage of the cspA transcript within the coding sequence and subsequent rapid removal of the fragments was also found in Rhodobacter capsulatus (a member of the alpha-proteobacteria), but no consensus cut sequence could be detected (Jäger et al., 2004). CspE was found to interfere with both the PNPase and RNase E of the degradosome machinery, inhibiting internal cleavage and removal of the poly(A) tails from mRNAs, thus stabilizing particular mRNAs (Feng et al., 2001). Bacteria without PNPase, which is a secondary cold shock protein, are cold sensitive (Clarke and Dowds, 1994; Goverde et al., 1998; Bae et al., 2000; Zangrossi et al., 2000). Curiously, this appears not to be true for Pseudomonas putida, indicating surprising differences between some species (Favaro and Deho, 2003). The coldtemperature induction of PNPase in E. coli occurs by reversal of its autoregulation. At 37°C, ribonuclease III cleaves the leader of the pnp mRNA, whereupon PNPase represses its own translation via unknown mechanisms. This latter step is inhibited after cold shock (Beran and Simons, 2001; Mathy et al., 2001). RNase H was recently found to be a cold shock protein, too. This protein, posttranscriptionally regulated by mRNA stabilization due to PNPase activity, subsequently regulates maturation of other mRNAs (especially small stable RNAs) by its exonuclease abilities. An RNase H mutant produces smaller colonies when grown at lower temperatures (Cairrão et al., 2003).

Other Cold-Inducible Proteins CspA induces and is part of the cold stimulon, directly or indirectly regulating 30 proteins, such as H-NS or GyrA (Madan Babu and Teichmann, 2003; Martinez-Antonio and Collado-Vides, 2003). H-NS is a histone-like nucleoid protein acting on DNA bending (La Teana et al., 1991; Brandi et al., 1994; Giangrossi et al., 2001b) and GyrA is part of topoisomerase II (Maxwell and Howells, 1999). Those promoters of secondary cold shock proteins contain one or more of the so-called “Y-box motif” CCAAT. Recognized by CspA, this Y-box motif subsequently activates transcription of the protein (Qoronfleh et al., 1992). This is true at least for H-NS, GyrA, and possibly other proteins (La Teana et al., 1991; Jones et al., 1992b). The enhanced level of GyrA together with H-NS, and HUβ increases the negative supercoiling of plasmids and chromosomal DNA (Goldstein and Drlica, 1984; Giangrossi

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CHAPTER 1.8

Heat shock Escherichia coli

37°C to 42–50°C Less negative SC

2 10

time (min)

σ

Topo I/Gyr HU + Gyr

–0.06 Negative SC Dnak

σ22

Cold shock Escherichia coli

2 –0.06

time (min)

60

37°C to 6°C

HU + Gyr Topo I (?)

σ Negative SC

Increased negative SC

Fig. 6. Effects of heat shock and cold shock on plasmid DNA. The left side of each panel shows the variation in the plasmid-specific linking difference (σ = δLk/Lko), dependent on the time of exposure to the shock temperature. SC, supercoiling; Topo, topoisomerase; Gyr, gyrase; and HU, a small, basic, heat-stable DNA-binding protein. From Lopez-Garcia and Forterre (1999).

et al., 2002). Why GyrB is not induced is unclear, but the induction of GyrA seems to be sufficient to increase the DNA twisting after cold shock. The DNA twisting itself regulates at least the induction of recA, the gene of another cold shock protein found in E. coli (Hulton et al., 1990; La Teana et al., 1991; Wang and Syvanen, 1992; Brandi et al., 1996; Hurme and Rhen, 1998). However, the increase in negative supercoiling was found to be transient after cold shock (Fig. 6). This shows that the open complex formation of transcription turned cold-insensitive after the adaptation of the entire system to low temperature (Krispin and Allmansberger, 1995; Lopez-Garcia and Forterre, 1999). In addition, most promoters of the –10/–35 type “close” in vitro below 15°C, which in turn may prevent protein synthesis for vital proteins below a certain threshold temperature (Minakhin and Severinov, 2003; Severinov and Darst, 1997). (Actually, the lowest temperature reported for growth in E. coli is 7°C; Kawamoto et al., 1989). Also, replication (Atlung and Hansen, 1999; Nyborg et al., 2000), protein folding (Seaton and

Vickery, 1994; Kandror and Goldberg, 1997; Vickery et al., 1997), and antitermination (Bae et al., 2000) are cold-regulated. A list of coldinducible proteins from E. coli is shown in Table 3. The list of cold shock induced genes has been extended recently by a genome-wide transcriptional analysis of cold shocked E. coli cells. New genes found by this study include transport or metabolism of diverse sugars and molecular chaperones (mopA, mopB, htpG, and ppiA). However, not all cold shock genes are displayed in this study (e.g. genes of the nusA-pnp operon), since some mRNA’s might be to unstable to be detected by this method (Phadtare and Inouye, 2004b). Some cold-inducible proteins from B. subtilis indicate that very different physiological processes such as chemotaxis (CheY), sugar uptake (Hpr), translation (ribosomal proteins S6, L7 and L12), protein folding (PPiB), and general metabolism (CysK, HvC, Gap and triosephosphate isomerase) are temperature regulated (Graumann et al., 1996b). A list of cold-stress induced proteins in B. subtilis described has been published by Graumann and Marahiel (1999b), and transcriptional profiling of the cold shock response in this organism was conducted by Beckering et al. (2002) and Kaan et al. (2002). Antitermination, which is mediated by CspA and other cold-shock induced Csps, was proposed to induce the genes of secondary coldinduced proteins (such as NusA, InfB, RbfA and Pnp) located in the region of the metY-rpsO operon. These Csps probably prevent secondary structure formation in the nascent RNA, which causes antitermination in ρ-independent terminator regions. The read-through produces a higher transcript level, which, in turn, increases the translation of such proteins (Bae et al., 2000; Zangrossi et al., 2000). nusA is an essential gene and NusA protein governs transcriptional elongation, pausing, termination and antitermination. The core RNA polymerase associates with the sigma factor (sigA) to form the holoenzyme that is capable of promoter recognition. As the polymerase complex enters the transcriptional elongation phase, NusA replaces SigA in the complex (Gopal et al., 2001). RbfA associates with the 30S subunit of the ribosome, enabling 16S rRNA maturation and interaction with mRNA (Xia et al., 2003). Another ribosome associated cold shock protein was discovered recently. Yfia is associated with the ribosomes in E. coli (as long as the growth is arrested) and disappears afterwards (Agafonov et al., 2001; Rak et al., 2002). A random observation of a cold-sensitive laboratory strain of E. coli led to the discovery of BipA. BipA was originally described as a protein induced after exposure to permeability-inducing

CHAPTER 1.8

Life at Low Temperatures

219

Table 3. Escherichia coli cold-inducible genes and their gene products. Gene aceE aceF

Product

Reference(s)

ahpC bipA (yihK) crhC csdA/dead cspA cspB cspG cspI des dnaA gyrA hns hscBA hupB infA infB infC lpxP nusA otsAB pnp

Pyruvate dehydro genase (lipomide) Pyruvate dehydro genase (dihydro lipoamide acetyltransferase) Alkyl hydroperoxidase reductase Ribosome associated GTPase RNA helicase (CrhC) Cold shock DEAD-box protein A Cold shock protein A Cold shock protein B Cold shock protein G Cold shock protein I Desaturase DNA A Gyrase subunit A H-NS (histone-like protein) Hsc66 (heat shock protein homolog) Nucleoid-assciated protein HUβ Initiation factor-1 Initiation factor-2 Initiation factor-3 Palmitoleoyl transferase NusA Trehalose synthesis Polynucleotide phosphorylase

rbfA recA rnr sodA tig ves yfia

Ribosome binding factor A RecA RNase H Superoxide dismutase Trigger factor TF Major cold shock protein family Ribosome-associated cold shock response protein

protein produced by neutrophils. The function of BipA at low temperature is not known (Pfennig and Flower, 2001). Another recent finding is the ves gene in E. coli, which is clearly cold inducible and shares some homology to cspH. But a mutant of this gene showed no phenotype at high or low temperature (Yamada et al., 2002). Organisms other than E. coli or B. subtilis may exhibit an “untypical” cold shock response. For example, in Listeria monocytogenes, a ferritin homolog was found under cold shock conditions and, similarly in Streptococcus thermophilus, an iron-binding protein being a member of the Dps family (Nicodeme, 2004 #2525; Hébraud, 2000 #2662). In Aeromonas hydrophila no CspA-like protein was found after cold shock, but only to transiently and weakly expressed 11 kDa proteins (Imbert and Gancel, 2004). The cold shock response is, as shown above, not a single event or a circumscribed response. Normally, cross-protection against other stresses is imprinted on the cells. A few recent reports on this finding include the induction of. barotolerance in Lactobacillus sanfranciscensis after cold stress (Scheyhing et al., 2004), NaCl tolerance in

Jones et al., 1987; Qoronfleh et al., 1992 VanBogelen and Neidhardt, 1990; Qoronfleh et al., 1992 Leblanc et al., 2003 Pfennig and Flower, 2001 Chamot et al., 1999 Jones et al., 1996 Goldstein et al., 1990 Etchegaray et al., 1996 Nakashima et al., 1996 Wang et al., 1999 Sakamoto et al., 1997b; Aguilar et al., 1998 Atlung and Hansen, 1999 Jones et al., 1992b La Teana et al., 1991; Brandi et al., 1994 Lelivelt and Kawula, 1995 Giangrossi et al., 2002 Giuliodori et al., 2004 Jones et al., 1987; Qoronfleh et al., 1992 Giuliodori et al., 2004 Carty et al., 1999 Jones et al., 1987; Qoronfleh et al., 1992 Kandror et al., 2002 Jones et al., 1987; Qoronfleh et al., 1992; Clarke and Dowds, 1994; Wang et al., 1996; Goverde et al., 1998 Dammel and Noller, 1995; Jones and Inouye, 1996 Jones et al., 1987; Qoronfleh et al., 1992 Cairrão et al., 2003 Smirnova et al., 2001b Kandror and Goldberg, 1997 Yamada et al., 2002 Agafonov et al., 1999; Agafonov et al., 2001

Shewanella putrefaciens (Leblanc et al., 2003), in Listeria monocytogenes sigB induction enhances freezing survival (Wemekamp-Kamphuis et al., 2004b), and survival of Vibrio parahaemolyticus after crystal violet challenge is higher after cold shock (Lin et al., 2004).

Cold Acclimation The term “cold acclimation” is used for cells that have adapted to low temperature after cold shock and have reached logarithmic growth with a new, now longer doubling time. In cold acclimated cells the internal processes have reached new equilibria, as could be shown for the protein content in Listeria monocytogenes (Liu et al., 2002), or the transcriptome in E. coli (Polissi et al., 2003). An interesting phenomenon related to cold acclimation is filamentation as exemplified by Salmonella, Escherichia or Pseudomonas strains kept at low temperature (Khan et al., 2003; Mattick et al., 2003a; Mattick et al., 2003b). This is also observed in Bacillus cereus and B. weihenstephanensis (K. Neuhaus and S. Scherer, personal observation). Whether filamentation of

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CHAPTER 1.8

bacteria grown under various stresses is a response to or just an aftereffect of the stress is not clear. However, a quadruple deletion of the cold inducible MCSP in E. coli shows filamentation already at 15°C (Xia et al., 2001b).

Cold Acclimation Proteins In contrast to cold shock protein (Csp) expression, cold acclimation protein (Cap) expression is at a higher level when the cold shock response has been downregulated (Hébraud and Potier, 1999). Not many reports deal with real Caps. However, both groups of cold-inducible proteins (Cips) are overlapping, since some cold-shock induced proteins have a transient maximum expression level but still show a higher level at low-temperature growth compared to growth at ambient temperatures. Our impression is, however, that the classifications given in the literature are not stringent. For instance, CspB is referred to as a cold shock protein in B. subtilis, but its expression is still higher at low than at ambient temperature (Weber and Marahiel, 2002; P. Graumann, personal communication; see the discussion section in Weber and Marahiel, 2002). According to Berger et al. (1997), CspB would be classified as a Cap. Therefore, the classification appears to be inconsistent and depends on the view of the particular researcher (Hébraud et al., 1994; Bayles et al., 1996; Michel et al., 1997; Mitta et al., 1997; Thieringer et al., 1998). Our simplified classification (Fig. 7) is based on Mitta et al. (1997) and Graumann and Marahiel (1999b) and the proposition that Cips are cold-induced proteins, irrespective of the kinetics of their regulation. Cips

can be divided into Csps with a transient peak of induction, regardless of the levels attained in comparison to pre-shock levels. The Csps may be divided into class I Csps (with an induction more than 10-fold) and class II Csps (with an induction less than 10-fold). Cold acclimation proteins (Caps) are not strongly cold shock induced (no clear peak), but levels steadily increase after a temperature downshift and remain higher (during the period of low-temperature cellular growth) than levels at ambient temperatures. In E. coli genes showing a higher transcriptional level 5 h after a cold shock to 15°C include genes encoding flagellar proteins, as well as the spermidine acetyltransferase speG (Phadtare and Inouye, 2004b). For Pseudomonas fragi, many different groups of Cips have been presented, but only four of them have been identified, and these belong to the CspA homology group. Two of them are designated “CapA” and “CapB,” and the other two (also heat shock induced) are designated “TapA” and “TapB” (temperature adaptation proteins; Hébraud et al., 1994; Michel, 1996). A further study dealing explicitly with cold-acclimation proteins names the CspA-homolog from Arthrobacter globiformis “CapA.” The protein level increases after cold shock and remains high for 24 h (Berger et al., 1997). In our opinion, the time period tested may be too short to determine whether this protein is a Csp or a Cap in this organism. When the cells were cold shocked from 25°C to 4°C, they had a subsequent lag phase of 14 h. For comparison, Yersinia enterocolitica, which can grow at –5°C like A. globiformis (Bergann et al., 1995), was cold shocked from 30°C to 10°C and had a lag of approximately 80 min. The level of the major Csps

Cold shock normal growth

lag

>10-fold

cold adapted growth

≤10-fold

bulk cap csp II

no peak cessation

csp I Fig. 7. Qualitative differences between different groups of cold induced proteins (Cips). Bulk proteins (brown) are also designated “housekeeping proteins.” After a cold shock, they decrease. Cold shock proteins (Csps) of group I (blue) have low concentrations at ambient temperature but are highly induced (>10-fold) after cold shock. They can remain on a higher or lower level during cold-adapted growth, as indicated by a split of the blue line. Csps of group II (green) are present at normal growth and are induced less than 10-fold. Cold acclimation proteins (Caps, pink) are induced at low temperature, but have no peak after a cold shock. Modified according to Thieringer et al. (1998).

CHAPTER 1.8

remains stable up to 3 h after cold shock. That means that the proteins are detectable at high level for more than twice as long as the lag phase (Neuhaus, 2000a). If this standard is applied to A. globiformis, the relatively large amounts of CspA should be detectable for 28 h, or even longer, before CspA finally disappears. A relatively recent list of cold-inducible proteins of B. subtilis and their classification is given by Graumann and Marahiel (1999b). They suggest several different proteins are Caps (e.g., L7/L12, L10, EFTs, EFTu, EF-G, Tig, GlnA, LeuC, ThrC, AroF and GuaB), which play different physiological roles in translation, protein-folding, and general metabolism. Recent studies suggest that in B. subtilis the transcription factor SigB, which is induced by cold shock, is important for continuous growth and sporulation at low temperature. Growth of a corresponding mutant is severely impaired at 15°C, but the mutant is rescued by addition of glycine betaine (Brigulla et al., 2003; Mendez et al., 2004). In Pantoea agglomerans, a 60-kDa protein was found to be a cryoprotective Cap; however, the nature of this protein is not known (Koda et al., 2001). A recent study with the psychrotolerant Listeria monocytogenes lists several genes as necessary for growth at 10°C. Most genes found were already known to participate in other stress responses, e.g., coldadaptation (flaA and flp), regulatory adaptive responses (rpoN, lhkA, yycJ, bglG, adaB and psr), general stress (groEL, clpP, clpB, flp and trxB), amino acid metabolism (hisJ, trpG, cysS and aroA), cell surface alterations (fbp, psr and flaA), and degradative metabolism (eutB, celD and mleA). Four further proteins with unknown function, only present in Listeria, were found, too (Liu et al., 2002). As can be seen from the diversity of the listed proteins found to be enhanced during growth at low temperature, many different aspects of the cell metabolism are affected to varying degrees. Interestingly, no (classical) MCSP was found, except the wellknown cold inducible flaA and flp genes (Bayles et al., 1996). For the identification of cold acclimation genes in Yersinia enterocolitica during prolonged growth in the cold, a genome-wide expression analysis was performed by creating random transcriptional fusions to the luxCDABE-reporter. This approach allowed the direct comparison of promoter activities (by comparing cell growth rates) of various genes at normal and low temperature. Out of 5700 investigated luxtransposon mutants, approximately 100 genes were detected with enhanced promoter activity at low temperature (compared to promoter activity under optimal growth temperature). Most of these genes could be placed into functional groups like motility proteins, transport

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221

proteins, and regulatory proteins (Bresolin et al., 2004). In any case, knowledge concerning cold acclimation proteins (which enable bacteria to grow constantly at low temperature) is limited, and further research in this area is needed. It would be especially interesting to examine psychrotolerant or psychrophilic organisms in contrast to their mesophilic counterparts. Such a comparison could be fruitful in genera containing psychrophile, mesophile, and thermophile species, such as Bacillus and others.

Compatible Solutes An emerging result from cold shock studies is that compatible solutes (such as glycine betaine, L-proline, and similar compounds) play also an important role in adaptation to the low temperature. However, how compatible solutes protect the cells against low temperature remains unclear. Several, possibly overlapping, scenarios are conceivable according to (Kandror et al., 2002): 1) compatible solutes may act as “chemical chaperones” against low temperature denaturation or aggregation. 2) Cold stress may cause oxidative stress, too. Some compatible solutes, e.g., trehalose, act as free radical scavengers. 3) Compatible solutes protect the membrane or 4) compatible solutes are induced in expectance of a possible drop below freezing. Listeria monocytogenes has at least two compatible solute transporters for glycine betaine: porter I is a Na/glycine betaine symporter and porter II is an ATP dependent transporter. Cold activated uptake of glycine betaine was most rapid between 7°C and 12°C (Mendum and Smith, 2002). Another compatible solute transporter is encoded by the opuC gene of Listeria, encoding for a carnitine transporter. This porter is also induced after chilling, and carnitine is also accumulated after osmotic or low temperature stress (Angelidis et al., 2002a), similar to observations made in B. subtilis by Brigulla et al. (2003). However, if the glycine betaine porter II is blocked, the increased carnitine uptake cannot completely restore the cryoprotective effect (Angelidis et al., 2002b). If all three compatible solute transporters are deleted, L. monocytogenes is severely impaired in growth at low temperature, but growth is not completely abolished (Wemekamp-Kamphuis et al., 2004a). In E. coli, the trehalose synthesis genes otsAB and the cryptic promoter (P1) of the proU transporter (important for mediating cytoplasmic accumulation of compatible solutes) are induced during low-temperature growth. Therefore, the cellular trehalose content increases up to eightfold after cold shock (Kandror et al., 2002; Rajkumari and

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Gowrishankar, 2001). If compatible solutes are added to the medium, L. monocytogenes and E. coli survive better in cold (Dykes and Moorhead, 2001; Shahjee et al., 2002). However, the situation might be different in psychrophilic bacteria, since (Mindock et al., 2001) have found no compatible solute accumulation in such Arthrobacter species at 4°C.

Bacterial Cold Sensors The reaction of bacteria towards cold stress should be as fast as possible. Currently, several processes that use a temperature signal to induce cellular processes have been suggested (for reviews, see Browse and Xin (2001) and Eriksson et al. (2002). Temperature is a factor that affects the whole bacterial cell immediately. Different mechanisms are used simultaneously to sense low temperature and these mechanisms might overlap as in the case of H-NS (see below). Note, furthermore, that many virulent bacteria have specially adapted temperature sensing mechanisms that determine whether they are inside or outside a host (for reviews, see DiRita et al., 2000; Eriksson et al., 2002; Gophna and Ron, 2003). A common threshold temperature for induction of virulence genes is around 30°C. Here, only the mechanisms involved in the cold shock or cold adaptation response (adaptation to temperature much lower than 30°C) will be discussed (see the section Pathogens in this Chapter).

DNA In bacteria, the degree of superhelicity of the DNA varies in response to changes in the ambient temperatures. In many examples, the expression of genes is dependent on DNA conformation (Eriksson et al., 2002). Supercoiling is mainly regulated in E. coli by topoisomerases I and II (Drlica, 1992; Hurme and Rhen, 1998; Tse-Dinh et al., 1997). But the conformation is fine tuned by proteins such as H-NS. H-NS appears to bind curved regions of DNA and is responsible for the cold repression of bacterial genes, possibly by denying open promoter complex formation necessary for transcription (Eriksson et al., 2002; Williams and Rimsky, 1997). Similarly, the promoter of the cold inducible histone-like protein HUβ is possibly affected by temperatures that stop transcription from the promoter sites P1 to P3, but not by temperatures that allow transcription from P4. The HUβ protein affects DNA structure, fine tuning transcription of many genes at low temperature (Giangrossi et al., 2002). A further analysis of the cold shock response on the DNA level is given by Golovlev (2003).

CHAPTER 1.8

An interesting facet is the involvement of the GATC methylation by the DNA methyltransferase Dam and its possible involvement in the cold shock response. According to this hypothesis, Dam is limited in fast growing cells (e.g. inside a host), leading to a hemi-methylated DNA which is more stable and displays a higher melting point. After shedding of E. coli into the environment, the cells experience a cold shock and the transcription of genes containing a GATC cluster will cease due to the high stability of hemi-methylated DNA. This effect might explain the decrease in transcription of some (many?) downregulated genes. However, this conclusions are only hypothetical so far (Riva et al., 2004).

Messenger RNA As has been discussed in the section The Cold Shock Response, in this Chapter, the mRNA of CspA is degraded rapidly at high temperature but is stabilized at low temperature. The cold sensor, therefore, is the folding characteristics and associated resistance to degradation of the mRNA. This reaction upon temperature downshift occurs rapidly, since the folding involves the mRNA that is already synthesized. Another example is the mRNA of σ32. At lower temperature, this mRNA is folded and therefore cannot be translated. At higher temperature, it is unfolded, becomes accessible to the ribosome, and the translated σ factor then switches on the heat shock response (Morita et al., 1999a; Morita et al., 1999b). A similar mechanism was described for σS, a stationary phase sigma factor. The transcription of rpoS, the gene for σS depends on dsrA, a small regulatory RNA which probably stabilizes the rpoS mRNA. The half life of dsrA is prolonged at 25°C compared to its half life at 37°C (Repoila and Gottesman, 2001). A similar system in which the mRNA acts as thermometer for thermal induction of a gene, is described by Chowdhury (2003) #2661.

Ribosome That the ribosome may act as a sensor for both heat and cold shock has been proposed (Fig. 8). After heat shock, the A-site of the ribosome is empty; at cold shock, the A-site is blocked owing to a stop in the initiation and translation of misfolded mRNA. Both situations lead to an increase or decrease, respectively, of the stringent response regulator, guanosine 5′triphosphate-3′-diphosphate and guanosine 5′-diphosphate-3′-diphosphate (collectively abbreviated [p]ppGpp). A decrease could be the signal for a cold shock response (VanBogelen and Neidhardt, 1990). For example, after a nutri-

CHAPTER 1.8

Life at Low Temperatures Cold shock

Heat shock

low concentration of charged tRNA

H-antibiotics kanamycin puromycin streptomycine

high - (p)ppGpp

223

high concentration of charged tRNA

empty A-site

blocked A-site

C-antibiotics chloramphenicol erythromycin tetracycline

Ribosome as sensor

H-state

Heat shock response

C-state

low - (p)ppGpp

Cold shock response

Fig. 8. Model of the ribosome as a sensor of temperature in bacteria. After a heat shock, translation proceeds faster than charged tRNA can be supplied, which may result in an empty A-site that is also affected by H-antibiotics. This could signal the ribosomal induction of the heat shock response (H-state) and increase of the guanosine 5′-triphosphate-3′-diphosphate and guanosine 5′-diphosphate-3′-diphosphate ([p]ppGpp) concentration. In contrast, cold shock leads to a reduced translational capacity of the cell and thereby a block of the A-site (as is also achieved by C-antibiotics) owing to a high concentration of charged tRNA. As a consequence, the cold shock response may be induced and the levels of (p)ppGpp are lowered, which in turn has been shown to increase the induction of cold-inducible proteins (Cips) after a cold shock. Redrawn from Graumann et al. (1996b).

tional downshift, the level of (p)ppGpp is increased and this leads to induction of DnaK and GroEL (both heat shock proteins; Schnier, 1987). Conversely, a nutritional upshift is coupled with a decrease in (p)ppGpp and leads to induction of CspA (Brandi et al., 1999a; Yamanaka and Inouye, 2001a). Additionally, a mutant lacking RelA ([p]ppGpp synthetase) and SpoT ([p]ppGpp hydrolase) is unable to produce (p)ppGpp and has a higher induction of cold shock proteins after a cold shock. This mutant phenotype seems to be preadapted to low temperature (Jones et al., 1992a).

Protein Changes in protein conformation, namely denaturation, are more pronounced after an increase in temperature. Such misfolded proteins are bound by chaperones, subsequently inducing a heat shock response (Arsene et al., 1999; Eriksson et al., 2002). However, conformational changes in proteins because of low temperature are also used for cold adaptation in some instances. An intriguing example of low temperature sensing by protein interaction is the aspartate chemotaxis of E. coli. The relevant thermosensors are transmembrane chemoreceptors or methyl-accepting chemotaxis proteins, one of

them (Tap) being a cold sensor (Nara et al., 1991). During adaptation, receptor methylation (catalyzed by the methyltransferase CheR) and demethylation (catalyzed by the methylesterase CheB) regulate the histidine kinase activity of the sensors. Thermosensing may be due to the specific temperature dependency of the methylation-demethylation equilibrium (Nara et al., 1996; Nishiyama et al., 1997; Nishiyama et al., 1999a; Nishiyama et al., 1999b). The action of H-NS (see the above section DNA) by influencing the conformation of DNA is itself to some extent dependent on the conformation of this protein. H-NS function is associated with oligomerization by means of a coiled-coil structure. This flexible structure stiffens at lower temperatures allowing better oligomerization and subsequent DNA binding of H-NS (Dorman et al., 1999; Smyth et al., 2000).

The Cytoplasmic Membrane Another mechanism of cold-temperature sensing involves the physical state of the membrane (for reviews, see Vigh et al., 1998; Sakamoto and Murata, 2002). For instance, it has been proposed that the thylakoid membrane acts as a cellular thermometer where thermal stress is sensed and transduced into a cellular signal leading to

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CHAPTER 1.8

37°C

25°C C

A DesK

DesK

Pi

B

DesK

Pi

P P

P P

+ Pi

DesR

DesR des

Des

desK

desR

–77 –28

des

–77 –28

desK

desR

ADP

ATP desKR mRNA

P

desKR mRNA

DesR DesR P

P

+1

des

desK

des mRNA

desR

desKR mRNA

Fig. 9. Model of des transcriptional control by two-component temperature signal transduction proteins. It is proposed that DesK assumes different signaling states in response to a temperature-induced change in membrane fluidity. This is accomplished by regulating the ratio of kinase to phosphatase activity such that a phosphatase-dominant state is present at 37°C, when membrane lipids are disordered (A), whereas a kinase-dominant state predominates upon an increase in the proportion of ordered membrane lipids after a temperature downshift to 25°C (B). DesK-mediated phosphorylation of DesR results in transcriptional activation of des (B) leading to synthesis of Des, which desaturates the acyl chains of membrane phospholipids (C). These newly synthesized unsaturated fatty acids inhibit des transcription either by favoring DesK dephosphorylation of DesR-P or by causing dissociation of DesR-P from its binding site (C). Adapted from Aguilar et al. (2001) and Cybulski et al. (2004).

the activation of heat shock (HS) genes (Horvath et al., 1998). In B. subtilis, a two-component signal transduction system was found involving a sensor kinase (DesK) and a response regulator (DesR). This system regulates the cold induction of the des gene coding for the D5-lipid desaturase. Unsaturated fatty acids (UFAs), which are the product of Des, act as negative signaling molecules of des transcription (Aguilar et al., 2001). Apparently, the physical state of the cytoplasmic membrane regulates the two-component system: DesK phosphorylates a DesR dimer after temperature downshift (which equals to ordered lipids in a more rigid membrane. The phosporylated DesR is able to bind to the promoter of des, inducing it thereby. After re-installing the fluid state of the membrane, DesK dephosphorylates DesR to inactivate it (Cybulski et al., 2004). The regulatory loop of DesK, DesR, Des and unsaturated fatty acids is shown in Fig. 9. In the cyanobacterium Synechocystis, two histidine kinases and a response regulator have been identified which regulate several genes at

cold temperature (Suzuki et al., 2000a; Suzuki et al., 2000b). Apparently, more than one coldresponding histidine kinase and probably several response regulators should be present in a cyanobacterial cell (Suzuki et al., 2001). Interestingly, the membrane bound histidine kinase senses not only cold but also osmotic stress with some overlap in the induced genes (Mikami et al., 2002). If the membranes are artificially rigidified by gene-engineering, some cold inducible genes are expressed at higher levels in this organism. However, some cold inducible genes do not respond to this intervention, suggesting that another cold sensor remains to be identified (Inaba et al., 2003). A similar two-component system is involved in upregulating phytopathogenic factors in Pseudomonas syringae. This system consists of the membrane-bound histidine protein kinase CorS and two transcriptional regulators, CorR and CorP, which induce virulence factors (Smirnova et al., 2002). Figure 11 summarizes the two-component sensor system from Synechocystis and B. subtilis.

CHAPTER 1.8

Life at Low Temperatures

Fig. 10. Osmostress-inducible and cold-inducible genes that were regulated by the sensor histidine kinase Hik33 in wildtype Synechocystis cells. Large and small circles enclose genes whose expression was induced by osmotic stress and cold stress, respectively. Rectangles in these circles enclose genes whose expression was regulated to a greater or lesser extent by Hik33 in cells under hyperosmotic stress and under cold stress. Genes outside rectangles appeared to be insensitive to the mutation in Hik33 in terms of their responses to the respective stresses. The rectangle in the overlapping region of the two circles encloses genes whose Hik33regulated expression was observed under both kinds of stress. From Mikami et al. (2002).

(a)

Osmostress

Hik33

Cold

Hik33 fabG murF hypA i infC rpl34

rpl28 groES

r

groEL groEL-2 clpB htrA imI ycf21 trpS hik31 hi k

lytB gifA

rh

Hik33 hliA hliB hliC igD crh rlpA




ggpS

n

Hik33 ycf39 livF

 d

rbpA rpl3 rpl11 rpl4 n

rn

rpoA

dr

gi


ili

(b) Baci

i sp. PCC 6803

225

Desk

HAMP

H

P

PAS H

H

HMG

P

P

D

Rer1 DesR ARNT P

HTH

D

Fig. 11. Schematic representation of the predicted structures of cold-sensing histidine kinases and signal-transducing response regulators involved in the regulation of expression of fatty acid desaturases in Synechocystis and B. subtilis. (a) Hik33 and Rer1 are the histidine kinase and response regulator, respectively, in Synechocystis sp. PCC 6803. In Hik33, the histidine kinase domain is indicated by pink rectangles, and the histidine residue possibly involved in the phosphorylation relay reaction is indicated by “H.” The HAMP (histidine-kinase-adenylyl-cyclase-methyl-binding protein phosphatase) domain and the PAS (PER-ARNT-SIM) domain (which is named after the period clock protein [PER] of Drosophila, the vertebrate aryl hydrocarbon receptor nuclear translocator [ARNT], and the single-minded protein [Whitby et al.] of Drosophila) are indicated by a blue diamond and a yellow circle, respectively. The PAS domain contributes to protein–protein interactions and, thus, Hik33 is likely to form a dimer, as shown in the figure. In Rer1, the receiver domain with a phosphorylatable aspartate residue is indicated by a green ellipse labeled “D.” The amino-terminal region is assumed to form a DNA-binding domain, which contains an HMG (high-mobility-group) box and the transcriptional activation domain of the aryl hydrocarbon receptor nuclear translocator (ARNT; (Suzuki et al., 2000a). (b) DesK and DesR are the histidine kinase and the response regulator, respectively, in B. subtilis. The DNA-binding domain is indicated by the HTH (helix-turn-helix) motif in the carboxy-terminal region that binds to the 5’upstream region of the des gene (Aguilar et al., 2001). In both systems, the cold-induced phosphorylation of histidine residues and the relay of phosphorylation to aspartate residues (indicated by arrows) have not yet been demonstrated. From Sakamoto and Murata (2002).

Cold Adaptation: General Remarks Mechanisms of cold adaptation in psychrotolerant and psychrotrophic bacteria remain poorly defined. Recent studies suggest multiple strategies to cope with low temperatures. A general conclusion drawn from such strategies is to allow more flexibility in any structures including mem-

branes, proteins or RNAs (Dalluge et al., 1996; Dalluge et al., 1997; Saunders et al., 2003). In membranes, unsaturated fatty acids are introduced to maintain ambient membrane fluidity. Proteins from psychrophilic bacteria are less rigid in structure owing to amino acid exchange (Gerday et al., 1997), or tend to dissociate easier into nonfunctional monomers because of a

S. Scherer and K. Neuhaus

weakening in hydrophobic bonds (Jahns and Kaltwasser, 1993; Ramstein et al., 2003).

Cold Adaptation of the Cytoplasmic Membrane General Strategies of Fatty Acid Alteration Membrane adaptation to different growth temperatures has been a target of research for a long time (de Mendoza and Cronan, 1983). There is now a large body of data dealing with the effect of temperature on the membrane composition of many species (for reviews, see Russell, 1997; Sakamoto and Murata, 2002). The lipid composition of the cytoplasmic membrane is of great importance for many cellular processes such as nutrient uptake, electron flow in respiration or photosynthesis, ATP synthesis, and others. A biological membrane is a highly complex and dynamic structure that can switch between different physical states. If bacteria are subjected to rapid chilling or freezing, a variety of damages can occur, like damage and release of lipopolysaccharides and altering the permeability of the membrane (Boziaris and Adams, 2001; Kempler and Ray, 1978; Ray and Speck, 1973; Riva et al., 2004). Besides having such mechanical effects, temperature influences the fluidity of the membrane. The lipid composition of the membrane, in combination with the temperature, controls the phase transition from the fluid phase to the semicrystalline or solid phase (Jones et al., 2002; for a review, see, e.g., Dowhan, 1997). To grow at low temperature, cells must have cytoplasmic membranes that retain sufficient fluidity to maintain a physical state supportive of the multiple functions of the membrane-a concept that has been termed “homoviscous adaptation” (Sinensky, 1974; Suutari and Laakso, 1994). Psychrophilic and mesophilic bacteria, as well as archaea, adjust the lipid composition of their membranes so that the proton permeability of their membranes remains within a narrow range. This phenomenon is termed “homeoproton permeability adaptation” (see Van de Vossenberg et al., 1995). The growth temperature-dependent alterations in fatty acyl chain composition are thus mainly aimed at maintaining the proton permeability of the cytoplasmic membrane at a rather constant level (Albers et al., 2000). For coldadapted bacteria such as Psychrobacter immobilis, this means that a decrease of the temperature would lead to a low proton permeability, which must be counteracted by an appropriate adaptation of the membrane lipids (Fig. 12). In general, several fatty acid changes are known to increase or decrease membrane fluidity in bacteria (for reviews, see Russell [1997]

CHAPTER 1.8

Proton permeability, k [s–1]

226

10–1

A

B

C

10–2

0

20

40

60

80

[°C]

Temperature Fig. 12. Graphic representation of the proton permeability of the psychrophilic bacterium Psychrobacter immobilis (line A), five mesophilic species represented by line B (Bacillus subtilis, Escherichia coli, Methanosarcina barkeri, Halobacterium salinarum and Halorubrum vacuolatum), and the hyperthermophilic Sulfolobus acidocaldarius, line C. The black squares represent measured proton permeabilities and the yellow area indicates the rather narrow range within which proton permeability is maintained and growth is possible. Note that some thermophilic and hyperthermophilic bacteria have higher proton permeability. See Albers et al. (2000), which is also the source of this figure. Table 4. Fatty acid changes influencing membrane fluidity in bacteria. Increase of fluidity Unsaturation Cis double bond Chain shortening Methyl branching Cis-unsaturation

Decrease of fluidity ↔ ↔ ↔ ↔ ↔

Saturation Trans double bond Chain lengthening Straight chain Straight chain

From Gounot and Russell (1999).

and Gounot and Russell [1999]). Most important for cold adaptation appears to be both unsaturation and chain shortening, but there are other adaptations (Table 4) which have been demonstrated experimentally. For instance, lpxP encodes a palmitoleoyl transferase. Palmitoleate is not present in E. coli grown at 30°C but comprises 11% of the fatty acid content in cells grown at 12°C. The lpxP gene was found to be 30-fold cold-inducible after 2 h. Thereafter, the activity gradually declines but does not disappear (Carty et al., 1999). A possible advantage suggested is that the palmitoleate content of the outer membrane provides a more effective barrier to harmful chemicals at low temperature (Vorachek-Warren et al., 2002).

Membranes in Psychrotolerants and Psychrophiles Recently, rather few reports have dealt with the membrane adaptation of psychrotolerant or psy-

CHAPTER 1.8

Life at Low Temperatures

chrophilic bacteria (Jones et al., 1997; Nichols et al., 1997; Whyte et al., 1999; Allen and Bartlett, 2000; Allen and Bartlett, 2002; Drouin et al., 2000; Edgcomb et al., 2000; Kumar et al., 2002). The psychrotolerant Listeria monocytogenes is a foodborne pathogen that can grow well at refrigeration temperature. Probably because of the medical importance of Listeria monocytogenes, some studies have been performed recently. When grown in continuous culture at 10°C in contrast to 30°C, this bacterium has a lower proportion of anteiso-C17:0 and a higher proportion of anteiso-C15:0 and short chain fatty acids (Jones et al., 1997). Similarly, Mastronicolis et al. (1998) found that cold shocked L. monocytogenes displayed increased anteiso-C15:0 in all lipid classes. Studies of fatty acid profiles of wildtype and cold-sensitive, branched-chain fatty aciddeficient mutants of L. monocytogenes suggest that the fatty acid 12-methyltetradecanoic acid (anteiso-C15:0) plays a critical role in lowtemperature growth of L. monocytogenes, presumably by maintaining membrane fluidity. The fluidity of isolated cytoplasmic membranes of the wildtype, and a cold-sensitive mutant of L. monocytogenes, grown with and without the

supplementation of 2-methylbutyric acid, has been studied (Annous et al., 1997; Edgcomb et al., 2000). These authors concluded that the fatty acid anteiso-C15:0 imparts an essential fluidity to the L. monocytogenes membrane and that this fluidity permits growth at refrigeration temperatures. However, even between closely related bacteria, differences in the membrane adaptation to low temperature can be found. The L. monocytogenes strains Scott A and CNL 895897 show differences in their pattern of branched fatty acids in response to low temperatures. The CNL strain uses, in addition to oddnumbered branched fatty acids found in both strains, substantial amounts of even-numbered branched fatty acids, too (Chihib et al., 2003). A similar example was reported from Sphingomonas, where one strain used unsaturated fatty acids and the other strain shifted from evenchain to odd-chain fatty acids (Männistö and Puhakka, 2001a). The production of increased proportions of membrane unsaturated fatty acids correlates with bacterial growth at low temperature or high pressure (Allen and Bartlett, 2002). Allen et al. (1999) characterized the fatty acids produced by

(Kdo)2

(Kdo)2 Growth at 30–42°C

O O HO P O OH O O

LpxL

Kdo2-Lipid IVA HO HO

OH HO O HO

O

HO O

O

O

HO O

O

O

O

HO

O O NH O P OH O OH HO

O

LauroylACP

OH

OH

HO

O

NH

O

227

LpxM

(Kdo)2

O 6’ O O 6 4’ 5’ O 4 5 O HO P O 1’ HO OH O 1 O 2’ NH O 3’ O 3 2 NHO P OH O O O OH HO HO O HO O O

MyristoylACP

O O HO P O OH O O HO

PagP

O

O

O

O

O

HO O

NH O

O

O

HO

O

O NH O P OH OH

HO O

PtdEtn

OH O O HO P O OH O O HO

O

O NH O

HO

HO O O

HO

O O

O NH O P OH OH

“Normal” Lipid A

HO

(Kdo)2

(Kdo)2

O

O HO P O OH O O

Cold Shock Palmitoleoyl12°C ACP LpxP

O

Lipid A under PhoP activating conditions

O O HO P O OH O O

O

O

HO O

NH O

O

O

HO

O NH O O P OH O OH HO

O

MyristoylACP LpxM

O

O O

O

O

O

O

HO O

NH O HO

O

NH O O P OH OH

HO

Cold Adapted Lipid A

Fig. 13. Biosynthesis of Kdo2 -lipid A during cold shock. In cells grown at 30°C or above, the key precursor Kdo2 -lipid IVA is utilized solely by the lauroyltransferase LpxL (Clementz et al., 1996). However, in cold-shocked cells, an additional acyltransferase, designated “LpxP” (Carty et al., 1999), is induced, which is proposed to incorporate palmitoleate at the same site normally reserved for laurate. In wildtype cells, the action of LpxL and LpxP is followed rapidly by the myristoyltransferase, LpxM, generating hexa-acylated lipid A (Brozek and Raetz, 1990; Clementz et al., 1997). About two-thirds of the hexa-acylated lipid A isolated from cells grown overnight at 12°C contains palmitoleate, and the remainder contains laurate. When the PhoP/PhoQ system is activated or when cells are grown on ammonium metavanadate, a portion of the lipid A molecules contain a palmitate residue at position 2, which is incorporated by the outer membrane enzyme PagP using glycerophospholipids as palmitate donors (Bishop et al., 2000). From Vorachek-Warren et al. (2002).

228

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the deep-sea bacterium Photobacterium profundum grown at various temperatures and pressures. In addition, oleic acid-auxotrophic mutants were isolated. One of these mutants, strain EA3, was deficient in the production of monounsaturated fatty acids and was both lowtemperature sensitive and high-pressure sensitive in the absence of exogenous 18:1 fatty acid. The authors conclude that monounsaturated but not polyunsaturated fatty acids are required for growth of P. profundum both at high pressure and low temperature. Note, however, that the fatty acid content does not always change dramatically in response to temperature. Könneke and Widdel (2003) examined a range of sulfate-reducing bacteria in their response of the fatty acid content (unsaturated vs. saturated). The highest levels of cisunsaturated fatty acids was measured in the psychrophilic species, but a substantial response in increasing amounts of unsaturated fatty acids

Desulfrigus oceanense

4˚C 10˚C

Desulfotalea psychrophia

4˚C 10˚C

Desulfotalea arctica

4˚C 16˚C

Desulforhophalus vacuolatus

4˚C 20˚C

Desulfobacter hydrogenophilus

4˚C 12˚C 20˚C 28˚C

Desulfobacter curvatus

12˚C 20˚C 28˚C

Desulfobacter tatus

12˚C 20˚C 28˚C

Desulfobacter postgater

12˚C 20˚C 28˚C

Desulfococcus multivorans

16˚C 28˚C

Desulfosarcina vanabiltis

16˚C 28˚C

Desulfovibrio desulfuricans

12˚C 20˚C 28˚C 60 80 100 0 20 40 Relative amount of unsaturated fatty acids (sum) among total fatty acids [%]

Fig. 14. Proportions of unsaturated fatty acids among total fatty acids in psychrophilic and mesophilic species of sulfatereducing bacteria grown at different temperatures. Analyses were carried out when cells were still growing and had reached three-quarters of the maximum (final) optical density. Note that only in Desulfobacter species a substantial increase in unsaturated fatty acids is visible. In all other species examined virtually no increase in unsaturated fatty acids can be found. Adapted from Könneke and Widdel (2003).

CHAPTER 1.8

at low temperature was only found in the genus Desulfobacter. All other genera (Desulfofaba, Desulfofrigus, Desulfotalea, Desulforhopalus, Desulfococcus, Desulfosarcina and Desulfovibrio) responded only with slight changes (Könneke and Widdel, 2003). A similar result was formerly reported about psychrotrophic Pseudomonas species (Bhakoo and Herbert, 1980).

Differences Between Closely Related Mesophilic and Psychrotolerant Strains Randomly selected strains of a bacterial collection of marine sea-ice bacteria from Antarctica were analyzed to obtain a profile of the membrane fatty acids. Results showed that shortchain saturated and unsaturated fatty acids were more common in the psychrotolerants when compared to psychrophiles. In contrast, branched-chain fatty acids were more abundant in the psychrophiles (Rotert et al., 1993). Such observations raise the question of whether differences in the capability of membrane adaptation to low temperature between closely related psychrotolerant and mesophilic strains (i.e., belonging to the same species) are responsible for the thermal type. Some species commonly harbor psychrotolerant as well as mesophilic strains. One example is Rhizobium leguminosarum, which is known as a mesophilic species growing poorly at temperatures under 10°C (Graham, 1992). However, psychrotolerant strains have been isolated from the Arctic legumes Astragalus and Oxytropis, and nitrogenase activity in Arctic nodules was detectable down to 0°C. The minimal and maximal growth temperature of isolates was 0°C and 27– 30°C, respectively (for a review on Arctic rhizobia, see Prévost et al., 1999). Psychrotolerant and mesophilic strains have also been isolated from the legume species Lathyus japonicus and L. pratensis (Drouin et al., 2000). These authors have determined the fatty acid profiles after growth at 25°C, at 5°C, and after cold shock from 25°C to 5°C. Interestingly, the degree of psychrotolerance of the strains did not correlate with their fatty acid composition. There is a vast body of literature concerning mesophilic and psychrotolerant isolates of the Bacillus cereus, which is a toxin producer (Granum and Lund, 1997; Dietrich et al., 1999; Prüß et al., 1999) in food (Mayr, 1999), and in soil (Von Stetten et al., 1999). All species of the Bacillus cereus group are so similar that they should be within the same species (Helgason et al., 2000). However, because members of the Bacillus cereus group are more or less medically important, placing them in a single taxon appears not to be sensible. Therefore, psychrotolerant

CHAPTER 1.8

isolates have been described as the new species Bacillus weihenstephanensis (Lechner et al., 1998). The difference in growth rate of the mesophilic and psychrotolerant strains is shown in Fig. 15. Very little is known about the physiological and genetic basis of cold adaptation of psychrotolerant strains of the Bacillus cereus group relative to mesophilic strains. We therefore analyzed the fatty acid composition (among other parameters) of a mesophilic Bacillus cereus and a very closely related psychrotolerant B. weihenstephanensis. Bacillus is known to have a branched-chain fatty acid profile (Table 5). Isoand anteiso-branched fatty acids are predominant, which is a characteristic observed in all species of Bacillus studied so far (e.g., Kämpfer, 1994). In both strains, iso-branched fatty acids increased about 6–7% at 12°C (in comparison to their amounts at 25°C) because of an increase of i-13:0 and i-16:0. A further increase of i-16:0 in the psychrotolerant strain at 7°C increased the level of branched iso fatty acids to nearly 50% of the total fatty acids. Upon lowering the temperature, straight-chain fatty acid and monounsaturated fatty acid levels decreased in response to changes in C16 fatty acid levels. These data confirm the hypothesis that bacilli adapt to decreasing environmental temperature by replacing the saturated straight-chain acids with the lower melting point branched-chain acids, or by changing to fatty acid branching instead of fatty acid unsaturation (Suutari and Laakso, 1992). Kaneda (1991) reported that mainly 12- and 13methyltetradecanoic acids (= a-15:0 and i-15:0) controlled the fluidity of membranes with branched-chain fatty acids. Indeed, these fatty acids constituted the major fraction in our study at 25°C but decreased at 12°C. At this latter temperature, an increase of i-13:0 was observed, which may also play a role in the control of membrane fluidity. In addition, we found some unidentified fatty acids in both strains. Especially, two fatty acids with retention times of 27 and 29 min increased in both strains to high levels upon lowering the temperature. We did not observe a correlation of the minimum growth temperature with the fatty acid composition. The nearly identical fatty acid pattern of the mesophilic and psychrotolerant B. cereus indicate that differences in lipiddependent membrane architecture may not be responsible for the substantially different growth rates of these strains at 12°C.

Carotenoids in Membranes Evidence is emerging that carotenoids in the cytoplasmic membrane also play a role in cold adaptation of some species (Jagannadham et al., 1996; Chattopadhyay et al., 1997). In vitro stud-

Life at Low Temperatures

229

ies with synthetic membranes of phosphatidylcholine demonstrated that the major pigments zeaxanthin, β-cryptoxanthin and β-carotene were bound to these membranes and decreased their fluidity (Jagannadham et al., 2000). In this respect it is interesting that Gram-positive bacteria collected from the Antarctic region show a predominance of pigmented isolates. In Arthrobacter agilis, collected from the Antarctic sea ice, pigmentation is due to a C-50 carotenoid induced at low temperature. Hypothetically, such carotenoids stabilize the membrane, since such C-50 carotenoids are only reported from other extremophiles and archea, coping with salt, cold and radiation stress (Fong et al. [2001] and references therein). This speculation fits the observation reported by Varkonyi et al. (2002) that some carotenoids are only low-temperature induced in the thylakoid membranes of the cyanobacterium Cylondrospermopsis raciborskii, possibly protecting them from reactive radicals. However, carotenoid-mediated stabilization of membranes and decrease in membrane fluidity seems to offset the increase in membrane fluidity accompanying fatty acid changes in low temperature habitats. Further research is needed to elucidate the interplay between carotenoids and fatty acids at low temperature and the exact role of the former.

Differences Between Thermotypes of Archaea as an Example A comparison of different thermotypes of microorganisms is possible by comparing single features, e.g., complete genomes or the MCSPs of different Bacillus species (Morra et al., 2003; Zeeb and Balbach, 2003a; Zeeb et al., 2003b; Zhou and Dong, 2003). Such an approach was chosen by Saunders et al. (2003), with interesting results for some Archaea: Comparative genomics between the two cold-adapted Methanogenium frigidum and Methanococcoides burtonii and other mesophile or (hyper-)thermophile Archaea revealed trends in amino acid and tRNA composition and structural features of proteins, which are to some extent applicable to eubacteria. Proteins from the cold-adapted Archaea are characterized by a higher content of noncharged polar amino acids, particularly Gln and Thr, and a lower content of hydrophobic amino acids, particularly Leu. Sequence data from nine methanogen genomes (optimal growth temperature 15–98°C) were used to generate 1111 modeled protein structures. Analysis of the models from the cold-adapted Archaea showed a strong tendency in the solvent-accessible area for more Gln, Thr, and hydrophobic residues and fewer charged residues. A cold shock domain (CSD) protein (CspA homolog) in M. frigidum,

230

S. Scherer and K. Neuhaus

CHAPTER 1.8 Growth ranges

1.5

Specific

A

growthrate

ot ole r

an ts

1.0

0.5

Me so ph iles

Ps yc hr

h–1

0

Climates

30 10 20 Temperature [˚C]

0

-10

Alpine II

1350

Psychrotolerants

98 96

av

86

e

80 45 40

e

era

ag

y av

ver

ag

er

al a

uar ge

500

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ly

nu

Jan

Temperature

Ratio of thermal types

Ju

An

2300

50

Temperature ranges

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Alpine I

40

0 0

Tropical

–10

7 10

0

20

30

40

50

Temperature [˚C] Annual average temperature in K–1 and proportion of psychrotolerants

0

Mesophiles

Spores + Vegetatives Spores

20 40 60 80 100 Proportion of psychrotolerants p [%]

D

Proportion 100 of psychro80 tolerant spores 60 p [%] 40 20 1/Tm

0 3.8

3.7

3.6

3.5

3.4

Temperature–1·103

3.3

3.2

3.1

[K–1]

Fig. 15. Difference in growth rate of a mesophilic Bacillus cereus and a psychrotolerant Bacillus weihenstephanensis strain, and influence of different climates on the ratio of mesophilic and psychrotolerant cspA genotypes. To obtain a holistic view, the diagrams have common axes. (B) and (C) share the climate axis, and (A), (B) and (D) share the temperature axes. Temperatures are displayed as both °C and K-1. A) Comparison between the growth ranges of psychrotolerant and mesophilic strains. B) Temperature ranges of a tropical climate, and a climatic sequence, consisting of one temperate climate at 500 m altitude and two temperate high-mountain climates at 1350 m and 2300 m altitude designated as alpine II and alpine I. January and July average temperatures (䊐) and annual average temperatures (䊉) are indicated. The white area indicates temperatures below 0°C, where no substantial growth occurs; the darker area marks the range between 0°C and 7°C, where only psychrotolerant strains grow well, and the brighter area highlights the growth range of the mesophilic strains. C) The ratios (in %) of psychrotolerant (dark bar) to mesophilic (bright bar) cspA genotypes are displayed for each climate. The upper bar of each pair gives this ratio for the total population, consisting of spores and vegetative cells; the lower bar shows the ratio for spores only. The 95% confidence intervals of the individual assays are indicated by error bars. The bold outer error bars of the temperate sample indicate the estimated intraclimatic mean variation. D) Proportion of psychrotolerant spores over the annual average temperature. This relation can be described by a tangens hyperbolicus function, with its point of inflection () shifted to 7°C. From Von Stetten et al. (1999).

CHAPTER 1.8

Life at Low Temperatures

231

Table 5. Major fatty acids of a mesophilic Bacillus cereus WSBC 10030 and a psychrotolerant Bacillus weihenstephanensis WSBC 10226 grown at different temperatures in percent. Mesophilic strain grown at

Psychrotolerant strain grown at

RTb,c

Fatty acid

12°C

25°C

7°C

12°C

25°C

12.95 16.37 19.82 23.32 26.48 29.86

i-12:0 i-13:0 i-14:0 i-15:0 i-16:0 i-17:0 Branched, iso a-13:0 a-15:0 a-16:0 Branched, anteiso 12:0 14:0 16:0 Straight, even 16:1 18:1 Unsaturated, even u.i. u.i.b u.i.b u.i.b Unidentified

0.9 13.4 4.9 16.6 7.0 2.4 45.2 3.9 7.2 1.6 13.7 0.4 3.7 7.8 11.9 7.5 1.4 8.9 1.4 8.7 3.7 0.7 14.5

1.0 8.5 5.6 18.1 2.0 3.1 38.3 3.7 10.2 4.1 20.2 0.8 5.6 10.4 16.8 10.3 1.5 11.8 1.2 4.9 2.2 0.7 9.0

1.6 14.7 3.8 7.5 18.0 0.8 46.4 6.2 6.5 0.0 13.2 1.9 4.4 2.9 9.2 4.3 1.8 6.1 1.9 10.1 7.7 1.1 20.8

2.1 15.3 4.1 10.1 6.8 2.5 40.9 8.8 8.9 1.6 20.7 1.8 4.3 5.5 11.6 8.4 1.0 9.4 1.0 5.7 5.3 1.2 13.2

1.4 8.9 3.4 13.3 2.6 5.1 34.7 5.3 9.0 3.6 20.9 1.4 4.9 8.1 14.4 11.6 1.7 13.3 1.4 3.3 4.9 1.4 11.0

16.64 23.60 26.63 14.19 21.12 27.88 27.38 33.35 25.71 27.00 29.00 29.27

The data in the column RT are retention times in minutes. All other numbers are percent of the individual fatty acids. The lines in italic reflect the sum of the indicated type of fatty acid and the lines below show the values of some specific fatty acids within each group. Abbreviations: WSBC, Weihenstephan Bacillus Collection, Microbial Ecology Group Weihenstephan; RT, retention time; and u.i., unidentified. a Thomas Kaplan and Siegfried Scherer, unpublished results. b Small peaks, representing less than 1% of total fatty acids are not listed. c See Byun et al. (2003).

two hypothetical proteins with CSD-folds in M. burtonii, and a unique winged helix DNAbinding domain protein in M. burtonii were identified. This suggests that these types of nucleic acid binding proteins play a critical role in cold-adapted Archaea. Structural analysis of tRNA sequences from the Archaea indicated that G+C content is the major factor influencing tRNA stability in hyperthermophiles but not in the psychrophiles, mesophiles or moderate thermo-philes. Below an optimal growth temperature of 60°C, the G+C content in tRNA was largely unchanged, indicating that any requirement for flexibility of tRNA in psychrophiles is mediated by other means. Recently, a proteomic determination of the cold adaptation in the Antartic archaeon, Methanococcoides burtonii has been undertaken. By this approach many proteins necessary for growth at low temperature were described, however, the function and interplay of these proteins

are still mostly unknown (Goodchild et al., 2004).

Response of Desaturases to Low Temperature Both anaerobic and aerobic mechanisms are responsible for the synthesis of unsaturated fatty acids (UFA) in bacteria. The anaerobic pathway, elucidated in detail for E. coli, produces cis-UFA by a specific 2,3-dehydrase acting at the C-10 level (for a review, see Cronan and Rock, 1996). A second mechanism is the introduction of double bonds into the fatty acids. The reaction is catalyzed by oxygen-dependent desaturation of the full-length fatty acid chain, either as an acyl-thioester or as a phospholipid fatty acid moiety, and requires a specific electron transport chain (see references in Aguilar et al., 1998).

S. Scherer and K. Neuhaus

Relative level of mRNA or protein (%)

35°C

CHAPTER 1.8

25°C

100

5

80

4

60

3

40

2

20

1

0

0 –6

0

6

12

18

Level of ω3-unsaturated fatty acids (mol %)

232

24

Time (h) Fig. 16. Changes of levels of desB mRNA (open circles), the encoded 3-desaturase (open triangles) and ω3-unsaturated fatty acids (closed diamonds) in Synechocystis after a temperature downshift from 35°C to 25°C. From Los and Murata (1999).

The molecular basis of the response of fatty acid adaptation to cold shock has been studied in some detail in unicellular cyanobacteria (Los et al., 1997; Sakamoto and Bryant, 1997a; Sakamoto et al., 1997b; Los and Murata, 1999). In cyanobacteria, four desaturase genes (desA– desD) have been reported; desA, B and D have been demonstrated to be cold-inducible in Synechocystis (cf. Fig. 16). In addition, desC mRNA has been reported to be upregulated within 15 min upon cold shock in Synechococcus. This upregulation appears to be due to an increased stability of des mRNA at low temperature. A series of mutants was generated by targeted mutagenesis of individual desaturases (Tasaka et al., 1996). Inactivation of desA plus desD in Synechocystis lead to a cold-sensitive phenotype that prevented this mutant from propagating at 20°C. Clearly, the desaturation of membrane lipids is an important factor in acclimation to low temperature. In contrast to cyanobacteria, B. subtilis has only a single desaturase gene, which was described mainly by two groups (Aguilar et al., 1998; Aguilar et al., 1999; Weber et al., 2001b). Cold shock induction of des occurs within 30 min and is almost exclusively controlled at the level of transcription, but unlike the situation in cyanobacteria, the stability of mRNA is not increased. Apparently, the des gene product is the only component of the B. subtilis desaturation system that is regulated by growth temperature. It is a typical transient cold shock induction, which would imply that desaturation

does not occur through de novo synthesis of fatty acids. Surprisingly, a des null mutant of B. subtilis has no phenotype even when cells are cold-shocked. However, this depends on the presence of isoleucine. In the absence of isoleucine, these mutants were cold sensitive. These data have been interpreted to mean that exogenous isoleucine triggers the switch from iso- to anteiso-branched saturated fatty acids, providing the organism with a second means to adapt membrane fluidity to low temperature (Klein et al., 1999; Fig. 17).

Adaptation of Protein Synthesis to Low Temperature Protein Synthesis and the Cold Shock Response The discovery by Broeze et al. (1978) that the initiation of mRNA transcription is impaired at low temperature indicated that the ribosome is a target of cold shock (Hurme and Rhen, 1998; Perrot et al., 2000). Acting as an RNA chaperone, CspA facilitates initiation and elongation of translation after cold shock (Jiang et al., 1997). Also, a ribosomal protein S21 homolog, which is encoded by rpsU, is cold induced in Sinorhizobium meliloti (O’Connell and Thomashow, 2000a) as well as in the cyanobacterium Anabaena variabilis (Sato, 1994; Sato et al., 1997). This protein may facilitate the binding of mRNA to the ribosome. Interestingly, rpsU is located downstream of cspA in S. meliloti. It may thus help the ribosome to function at low temperatures in the same way as other cold shock proteins (such as RbfA; Jones and Inouye, 1996; Huang et al., 2003). Possibly, small cold shock proteins (e.g., CspA), which appear to be synthesized continuously in some organisms (Graumann et al., 1997; Yamanaka et al., 1999b), may help render the ribosomes able to participate in translation at cold temperatures (i.e., transform them into cold-insensitive ribosomes), but this has not been demonstrated experimentally until now. In any case, by tagging CspB in B. subtilis with the green fluorescent protein, and ribosomal protein L1 with the blue fluorescent protein, CspB and ribosomes were seen to colocalize in the cell (Mascarenhas et al., 2001; Weber et al., 2001c). The level of inactive ribosomes determines the extent of the expression of the cold shock response. Once a balanced translational capacity is achieved, the cold shock response is repressed. At least four proteins (RbfA, initiation factor [IF] 2, CsdA/DeaD, and pY/Yfia) have been proposed as mediators the ribosome’s transformation into a cold-insensitive state (Jones and

CHAPTER 1.8

valine

a

α -keto-iso-valerate b

iso-butyryl-CoA

Life at Low Temperatures

leucine

a

α -keto-iso-caproate b

iso-valeryl-CoA

233

isoleucine

a

keto- β -methyl-valerate

2-methyl-butyrate

b

2-methyl-butyryl-CoA

fatty acid biosynthesis

iso C14:0 iso C16:0

iso C15:0 iso C17:0

anteiso C15:0 anteiso C17:0

Fig. 17. Schematic representing branched chain fatty acid biosynthesis in Bacillus subtilis and its dependence on external supply of valine, leucine and isoleucine. The isoleucine-based pathway offers one possible avenue for membrane adaptation to low temperatures. From Klein et al. (1999).

Inouye, 1996). The ribosomal binding factor A (RbfA) was found to be a suppressor of a coldsensitive mutation in the 16S rRNA. Cells lacking RbfA exhibit a cold-sensitive phenotype (Dammel and Noller, 1993; Dammel and Noller, 1995), perhaps because the 16S RNA is not processed properly (Bylund et al., 1998). CsdA was found in a 2D-gel analysis of 70S ribosomes from cold-shocked E. coli and designated “cold shock DEAD-box protein A.” This protein is a homolog of the DEAD-box helicases and possesses RNA unwinding activity. Again, a CsdA mutant is impaired in growth at low temperatures and has the cold-sensitive phenotype of elongated cells (Jones et al., 1996). Recently, it was reported that CsdA is involved in biogenesis of 50S ribosomal subunits. Presumably the RNA helicase activity of CsdA permits a structural rearrangement during 50S biogenesis at low temperature (Charollais et al., 2004). Finally, IF2 is needed for initiation of mRNA translation at the

ribosome (Moreno et al., 2000). New data about a ribosome modification after cold shock were published by Agafonov et al. (2001). Ribosomes of cold shocked E. coli are shown to be associated with a protein called “PY” or “Yfia” (Rak et al., 2002). However, this protein apparently disappears when the growth arrest is resolved (Kalinin et al., 2002). PY blocks the P as well as the A site of the ribosome, inhibiting translation initiation during cold shock but not under normal growth conditions. Only cold shock genes such as cspA may be able to override PY inhibition. By blocking the translation of all but cold shock proteins, the cell diverts all translation factors to the synthesis of cold shock proteins, thus ensuring its survival in the cold (Vila-Sanjurjo et al., 2004). This finding might explain the initiation inhibition after cold shock, originally observed by (Broeze et al., 1978). O’Connell et al. (2000b) screened cold-shock gene loci in Sinorhizobium meliloti by using a

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luxAB reporter transposon. Unexpectedly, they found that the transposon of many coldinducible mutants was inserted in the 16S and 23S rRNA genes. Subsequent experiments confirmed that transcription of all three rrn operons of this bacterium is induced by cold shock. Since the number of ribosomes is usually positively correlated with growth rate, one would expect ribosome synthesis inhibition when growth at low temperature is downregulated. However, the cell may upregulate ribosome synthesis because protein synthesis is severely inhibited at low temperature but is needed for survival.

Cold Adaptation of the Ribosome As has been described above, protein synthesis of mesophilic bacteria is a target of the cold shock response. For protein synthesis, a proper function of tRNA is essential. Many posttranscriptional modifications of tRNA are known. The study of three psychrophilic bacteria from the genera Moritella and Vibrio revealed that, among other posttranslational modifications, these organisms contained 40–70% more dihydrouridine than did mesophilic bacteria (Dalluge et al., 1997). Nuclear magnetic resonance studies showed that dihydrouridine leads to a higher local flexibility of RNA molecules (Dalluge et al., 1996). Apparently, therefore, the role of the elevated content of this modified nucleoside is to increase local conformational flexibility of tRNA under low temperature conditions where thermal motions and intermolecular interactions of biomolecules are compromised. Interestingly, downstream of the cold inducible rbfA gene, a pseudouridine tRNA synthase gene (trueB) is located in E. coli (S. Scherer and K. Neuhaus, unpublished observations). In 1969, ribosomes prepared by Nash and Grant (1969) from a psychrophilic Candida gelida were inactivated rapidly at 40°C, whereas the ribosomes from a mesophilic Candida utilis were unaffected by a similar treatment. Ribosomes prepared by Szer (1970) from a psychrophilic Pseudomonas were functional at 0°C and contained a factor which could be washed off, leaving the ribosomes functional at 25–37°C but not at lower temperature. The ribosomes of the mesophile E. coli became activated at low temperature upon addition of this factor. Both authors thus concluded that the ribosomes of cold-adapted microorganisms should be structurally different from mesophilic ones, but the identity of the proteins involved is still unknown. Cold-adapted microorganisms may therefore have structurally different ribosomes when compared to mesophilic bacteria. The comparison of mesophilic and psychrotolerant isolates from the B. cereus group showed a systematic difference

CHAPTER 1.8

in the structure of 16S rRNA (Lechner et al., 1998; Von Stetten et al., 1998). Interestingly, both signatures systematically contain A or T in psychrotolerant strains, and G or C in mesophilic strains. One may therefore speculate that the flexibility of the ribosome at low temperature may be increased in some parts of the molecule in the psychrotolerant isolates. However, the occurrence of specific sequence motifs in psychrotolerant strains is not necessarily due to a positive selection pressure associated with this ribosome’s function but could be a consequence of neutral drift processes. Therefore, further analysis of the genomic DNA from a wide range of isolates was undertaken (Prüß et al., 1999). This analysis showed that B. cereus group strains have between 6 and 10 copies of 16S rDNA. Moreover, a number of these environmental strains have both rDNA operons with psychrotolerant signatures and rDNA operons with mesophilic signatures. The ability of these isolates to grow at low temperatures correlates with the prevalence of rDNA operons having psychrotolerant signatures, indicating specific nucleotides within the 16S rRNA play a role in psychrotolerance (Fig. 18). In vivo measurement of protein synthesis in a psychrotolerant B. weihenstephanensis and a mesophilic B. cereus clearly showed that 35S-methionine incorporation at low temperature occurs faster by a factor of four (T. Kaplan et al., unpublished data).

Protein Structure and Enzyme Activity In general, the temperature optima of enzymes from cold-adapted bacteria have been reported to be well above the growth optimum (e.g., Reichhardt, 1998), but those of extracellular enzymes from Arctic and sea ice bacteria have been reported to be as low as 15–20°C (Huston et al., 2000). Usually, the enzyme activity at low temperature is comparatively high (Sun et al., 1998) and the thermostability of cold adapted enzymes is reduced significantly. An example of temperature-dependent activity of the same enzyme isolated from a psychrophilic, mesophilic and thermophilic bacterium is shown in Fig. 19. However, investigation of the molecular basis of cold-active enzymes from psychrophiles has only recently received increased research attention owing to novel opportunities for biotechnological exploitation (Russell, 1998). The application of these enzymes offers considerable potential to the biotechnology industry, for example, in the detergent and food industries, for the production of fine chemicals, and in bioremediation processes (Gerday et al., 2000).

CHAPTER 1.8

Life at Low Temperatures

A

B 0.11

0.07 0.05 0.03

2.5

1.4

Growth rate (gen/h)

Growth rate (gen/h)

Growth rate (gen/h)

C

1.6

0.09

235

1.2 1 0.8 0.6 0.4

2 1.5 1 0.5

0.2

0.01

0 –0.01 0

0 0

50 100 Psychrotolerance (%)

50

100

0

Psychrotolerance (%)

50

100

Psychrotolerance (%)

Fig. 18. Comparison of growth rates and psychrotolerance indices. Cultures were grown at 10°C (A), 28°C (B), and 42°C (C). The growth rates are plotted against the percentage of psychrotolerant signatures. A psychrotolerance index of 50% means that one half of the operons of this strain carries the psychrotolerant signature and the other half carries the mesophilic signature. The experiment was done two to six times, and the mean of the populations were determined. Data are from Prüß et al. (1999).

Chemical reactions are characterized by a strong dependency of the reaction velocity on the reaction temperature. The decrease of the rate constant can be described by the Svante Arrhenius equation K = A e–Ea/RT where R and T represent the molar gas constant and absolute temperature, respectively, and A is the frequency factor. Typically, a decrease of the reaction temperature by 10°C will lead to a decrease of the reaction rate by a factor of 1.5– 4 (Q10 value). Notably, the greater the activation energy Ea, the stronger is the temperature dependency of the reaction rate (the reaction rate constant is K). Reactions with low activation energies will only slightly depend on the reaction temperature.

A

Escherichia coli

Bacillus

100

1000

80

800

kcat (s–1)

β galactosidase activity (%)

Arthrobacter

The influence of the reaction temperature on the reaction rate is more complicated when enzyme-catalyzed reactions are considered (for reviews, see Gerday et al., 1999; Lonhienne et al., 2000; Feller, 2003a; Lonhienne et al., 2000). In this case, substrate concentration, enzyme concentration as well as the enzyme-substrate interaction play an important role. At nonsaturating substrate concentration, the reaction velocity depends also on the Km, which is influenced by the nature of the interaction of the enzyme with the substrate. An electrostatic interaction will be weakened by an increase in temperature, while the hydrophobic interactions tend to be stabilized. Therefore, the reaction velocity of enzymes will be differentially influenced by temperature because of the relative contribution of electrostatic versus hydrophobic forces. Such factors

60 40

600 400 200 0

20

0 B

0 0

15

30 45 60 temperature (°C)

20 40 60 Temperature (°C)

80

75

Fig. 19. Graphic comparison of the thermodependence of enzymes. A) β-Galactosidase from the psychrophile Arthrobacter D2 (blue), the mesophile Escherichia coli (violet) and a thermophilic Bacillus (red). Adapted from Brenchley (1996). B) αAmylase from a psychrophilic (䊉) and a mesophilic (䊊) organism.

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CHAPTER 1.8

have to be considered to understand the cold adaptation of enzymes. Generally, one would assume that enzyme stability, flexibility, and activity have to be properly adjusted to the low temperature. Another aspect of proteins from psychrotrophic organisms is not only the increased thermolability of the protein itself, but also the increased dissociation of monomers or heterodimers. As examples the HNS-like protein from the psychrophilic Psychrobacter should be mentioned. The α-helical domain of this protein displays weaker intermolecular interactions, which may account for the low thermal stability at 37°C (Tendeng et al., 2003). In E. coli, H-NS has two isoforms, HUα and the cold-inducible isoform HUβ. HUβhomodimers show weaker intermolecular interactions (Ramstein et al., 2003). Recently, many cold active enzymes have been purified and characterized, both at the biochemical and structural level. This section does not cover the biochemical and biophysical aspects;

instead, the reader is referred to recent reviews (Gerday et al., 2000; Russell, 2000; Lonhienne et al., 2000; D’Amico et al., 2002a; Feller, 2003a; Feller and Gerday, 2003b) or to Table 6, which lists recent studies of psychrotrophic enzymes. In summary, several adaptations to low temperature are found in different enzymes from psychrophilic bacteria (Table 7). Notably, in no case have all of these adaptations been realized in one protein; each protein has a couple of such changes which is sufficient to render the enzyme cold active. The rules governing their adaptation to cold appear to be relatively diverse, and they are only beginning to be understood. Due to the widespread occurrence in all thermal types and the relative small size, CspB from Bacillus species is a favored model protein for examining thermostabilization, folding and similar structural effects. The findings obtained on CspB might be generalized on other protein families occuring in different thermal types of bacteria, but are beyond the scope of this review. The

Table 6. Recent publications dealing with cold adapted enzymes from psychrotrophic organisms. Enzyme characterized

Species

Reference(s)

3-Isopropylmalate dehydrogenase adenylate kinase alcohol dehydrogenase Alkaline phosphatase Alkaline phosphatase aminopeptidase Chitinase Chitobiase Citrate synthase Dihydrofolate reductase Esterase Family 8 xylanase Glutamate dehydrogenase hydrolytic enzymes Isocitrate lyase

Vibrio sp. I5 Bacillus globisporus Moraxella sp. TAE123 Shewanella sp. Vibrio sp. AP Colwellia psychrerythraea strain 34H Alteromonas sp. O-7 Arthrobacter sp. TAD1 Arthrobacter sp. DS2-3R Moritella profunda Psychrobacter sp. Ant300 Pseudoalteromonas planktis Psychrobacter sp. TAD1 divers Colwellia maris

Lipase L-Threonine dehydrogenase malate dehydrogenase Malate synthase NAD+ dependent dehydrogenases Omithine carbamoyltransferase Pectate lyase peptidyl-prolyl cis-trans isomerase proteases Protein-tyrosine phosphatase Replication protein Rep Serine hydrolase Serine peptidase Subtilisin-like serine protease α-Amylase β-Galactosidase β-Galactosidase β-Galactosidase β-Galactosidase

Pseudomonas fragi Cytophaga sp. KUC-1 Moritella sp. strain 5710 Colwellia maris Shewanella PA-43 Moritella abyssi Pseudoalteromonas haloplanktis Shewanella sp. SIB1 Pseudomonas sp. Shewanella sp. Psychrobacter sp. TA144 Acinetobacter sp. No. 6 Shewanella sp. PA-43 Vibrio sp. PA44 Pseudoalteromonas haloplanktis Arthrobacter psychrolactophilus Arthrobacter sp. SB Pseudoalteromonas haloplanktis Pseudoalteromonas sp. TAE 79b

Svingor et al., 2001 Bae and Phillips, 2004 Liang et al., 2004 Murakawa et al., 2002 Asgeirsson and Andresson, 2001 Huston et al., 2004 Orikoshi et al., 2003 Lonhienne et al., 2001 Gerike et al., 2001; Kumar and Nussinov, 2004 Xu et al., 2003b Kulakova et al., 2004 Van Petegem et al., 2003 Camardella et al., 2002 Groudieva et al., 2004 Watanabe et al., 2001; Yoneta et al., 2004; Watanabe and Takada, 2004 Alquati et al., 2002 Kazuoka et al., 2003 Saito and Nakayama, 2004 Watanabe et al., 2001 Irwin et al., 2001b Xu et al., 2003a Truong et al., 2001 Suzuki et al., 2004 Vazquez et al., 2004 Tsuruta et al., 2004 Duilio et al., 2001 Suzuki et al., 2002 Irwin et al., 2001a Arnorsdottir et al., 2002 Claverie et al., 2003; D’Amico et al., 2002b Nakagawa et al., 2003 Hoyoux et al., 2001 Fernandes et al., 2002 Coker et al., 2003

CHAPTER 1.8 Table 7. Adaptation of cold active enzymes in comparison to their mesophilic counterparts. – More polar and less hydrophobic residues – Additional glycine residues and low arginine/lysine ratio – Fewer hydrogen bonds, aromatic interactions, and ion pairs – Lack of or fewer salt bridges – Additional or extended surface loop(s) with increased polar residues, or decreased proline content (improves solvent interactions), or both – Modified alpha helix dipole interactions – Reduced hydrophobic interactions between subunits – Weaker calcium binding From Russell (2000) and Arnorsdottir et al. (2002).

reader may consult Garcia-Mira et al. (2004), Makhatadze et al. (2004), Garofoli et al. (2004), and similar publications.

Metabolic Activity and Growth Bacteria experiencing a cold shock normally adapt by induction of the cold shock response and cold shock acclimation proteins. If they are kept at temperatures below the minimal growth temperature, cells tend to die over time. Estuarine and marine Vibrio species seem to disappear under low temperature conditions (e.g., below 15°C) from their habitat but reappear with increasing temperatures. Such organisms enter a so called viable but not culturable (VBNC) state. During this VBNC state, the cells become coccoid, whereas normally they are rod shaped and their metabolic activity is maintained. A resuscitation is possible by shifting the culture to higher temperature (e.g., to 37°C for 24 h) before plating (Carroll et al., 2001; Datta and Bhadra, 2003; Johnston and Brown, 2002). Other organisms are also known to enter a VBNC state after exposure to low temperature, as e.g., Aeromonas hydrophila (Mary et al., 2002). The latter strain was reported not to have a Csp similar to CspA of E. coli, which might contribute to the entering of a VBNC state (Imbert and Gancel, 2004). The VBNC condition might allow such organisms to become resistant and dormant below temperatures permissive for their growth and survive with a minimal metabolic rate. The lowest temperatures at which metabolically active bacterial communities exist has been reported to be –12°C to –17°C (Carpenter et al., 2000).

Motility In any environment, bacteria use motility to either find nutrients or associate with a surface. Whether bacteria move in subzero environments, such as sea ice, is unclear since the lowest temperature tested for motility or chemotaxis is

Life at Low Temperatures

237

5°C. Though Colwellia psychrerythraea was shown to be motile at –10°C, the minimum temperature for its growth is reported as –5°C. The swimming speed dropped to 28 µm/s before ceasing (Junge et al., 2003).

Nutrient Uptake Nutrient uptake is a basic prerequisite for growth. Algae as well as bacteria have a reduced affinity for nitrate at low temperature (Nedwell and Rutter, 1994), and on the basis of the different nitrate uptake and ammonium uptake responses to temperature, dependency on ammonium as an inorganic nitrogen source is suggested to increase at low temperatures (Ray et al., 1999). Apparently, mesophilic bacteria have an enhanced substrate requirement at minimal growth temperatures (Wiebe et al., 1992). Therefore, one would expect that high activity of transport systems at low temperatures is a prime target of cold adaptation of psychrophilic bacteria (Russell, 1990a). Some reports state that sugar transport is largely independent of temperature in psychrophilic yeast and psychrotolerant bacteria (for a review, see Herbert, 1986). To our knowledge, only one transport system from a psychrophilic bacterium has been studied so far. The 14CH3NH3+ uptake activity of a psychrophilic marine bacterium Vibrio sp. was markedly higher at 0–15°C, and the apparent Km value for the uptake of 14CH3NH3+ did not change significantly over the temperature range 0–25°C. Thus, the NH4+ transport system of this bacterium was highly active at low temperatures (Chou et al., 1999). Assessment of the temperature dependency of this system (Fig. 20) demonstrated its unusual psychrophilic properties.

Carbon Metabolism and Electron Flow Carbon metabolism and electron flow is also affected by temperature. In chilling-sensitive cold-stressed plants, a decrease in temperature inhibits respiration, but not much is known about this response in bacteria. Cold stress seems to induce changes in the carbon flow of a given organism, either by increasing cold sensitive key enzymes necessary for certain metabolic pathways or by switching to alternative pathways or cold adapted isoenzymes. Cold stress induces a change from respiratory metabolism to anaerobic lactate formation in psychrotrophic Rhizobium strains (Sardesai and Babu, 2000). Analysis of specific activities of glucose-6-phosphate dehydrogenase and 6phosphogluconate dehydrogenase of the pentose phosphate pathway showed the upward regulation of alternate pathways of carbohydrate metabolism under cold stress, resulting in rapidly

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CHAPTER 1.8

14CH3NH + 3

uptake rate (nmol/min/mg protein)

0.8

0.6

0.4

0.2

0 0

10

20

30

40

Temperature (°C) Fig. 20. Temperature dependence of the cold-adapted ammonium uptake system of the psychrophilic marine bacterium Vibrio sp. strain ABE-1. From Chou et al. (1999).

generated energy to overcome the stress. The glycolytic activity was also markedly stimulated by a factor of 2.5 in Lactococcus lactis upon cold shock from 30°C to 10°C (Wouters et al., 2000a). Upward regulation of malate dehydrogenase suggests that it is a critical input for cold tolerance (Sardesai and Babu, 2001). The cold-stress mediated decrease in the poly-β-hydroxybutyrate (PHB) in a psychrotolerant Rhizobium was due to an inhibition of PHB synthesis rather than an increase in its breakdown (Sardesai and Babu, 2001). A downshift in temperature had marked effects on carbon and electron flow in a methanogenic archaeal community in rice field soil, leading to a dominance of psychrotolerant homoacetogenesis (Fey and Conrad, 2000). Glucose oxidation was also found to be temperature-regulated. At low growth temperature, Pseudomonas fluorescens accumulated 2-ketogluconate in the medium as the major oxidation product of glucose. At 30°C, no 2-ketogluconate was excreted at any time (Lynch et al., 1975a; Lynch et al., 1975b). Also, a marked effect of temperature on diauxic growth with glucose and organic acids was observed in this bacterium. Organic acids were preferentially used at 30°C during the first growth phase, and glucose utilization was delayed until onset of the second growth phase. At 5°C, glucose utilization was not repressed during the first growth phase (Lynch and Franklin, 1978). Another psychrotrophic member of this genus, Pseudomonas syringae,

upregulates urocanase for histidine utilization upon a temperature downshift (Janiyani and Ray, 2002). To cope with cold, psychrophilic Colwellia maris expresses a thermolabile isocitrate lyase. This isocitrate lyase is able to utilize its substrate at lower temperatures because of a lower temperature optimum. The same organism has two isocitrate dehydrogenase isoenzymes: one with mesophilic (IDH-I) and the other with psychrophilic characteristics (Ochiai et al., 1979). Accordingly, both the tricarboxylic acid and glyoxylate cycles are important for growth in cold (Watanabe et al., 2002). A lipase produced by a psychrotrophic Pseudomonas strain was found to have the lowest temperature optimum of 35°C in vitro, to have higher activity at low temperature, and to be thermolable compared to other lipases from the same enzyme subfamily (Rashid et al., 2001). The authors conclude that this lipase has adapted to function within the growth range of its host (i.e., –5°C to 35°C). The psychrotrophic Acinetobacter sp. HH1-1 undergoes several metabolic changes in adaptation of its carbon metabolism if exposed to low temperature: 1) Isocitrate lyase is mainly found in the culture supernatant at low temperature. Whether this is due to leakage, as the authors of the study suggested, or to active transport is not clear. 2) The cell associated esterase activity increases and seems to be important for growth at low temperature. 3) Extracellular lipolytic enzymes and production of extracellular polysaccharide are negatively affected at lower temperatures (Barbaro et al., 2001).

Growth Rates Many more physiological processes are adapted to low temperature in psychrophilic microorganisms. Examples are histidine utilization (Kannan et al., 1998), sulfate reduction (Knoblauch and Jørgensen, 1999a; Knoblauch et al., 1999b), transcription (Ray et al., 1999; Uma et al., 1999), adaptation of the outer membrane of Gramnegative bacteria (Ray et al., 1994; De et al., 1997), reduction of the polar polysaccharide capsular layer (Mindock et al., 2001), carotenoid synthesis (Chattopadhyay et al., 1997), or exoenzyme secretion. The latter is even maximal at –2°C to +4°C in four psychrophilic Antarctic bacterial strains (Feller et al., 1994). In toto, numerous cold-adapted physiological reactions contribute to, and determine, the growth rate of cold-adapted bacteria (see also the section The Cold Shock Response and Cold Adaptation in this Chapter). As a result, at low temperature, growth rate is higher than in mesophiles, and the lower limit is lower. In principle, the lower growth limit is determined by the

CHAPTER 1.8

Life at Low Temperatures

Brenchley (1996) noted that only a few studies have been reported on growth rates at low temperatures. Table 8 lists some doubling times at low temperatures that have been compiled from Temperature (°C) 30

17

0

0 b −1

Ln k

freezing temperature of the cytosol. Most cells remain unfrozen at –10°C to –15°C because of the physical properties of the aqueous solvent systems inside and outside the cells (see Russell, 1990a). These physical boundaries thus determine the absolute lower growth temperature limit in general. The deepest temperature of metabolic activity in bacteria has been reported as –17°C (Carpenter et al., 2000). The maximal specific growth rate of a psychrotolerant Pseudomonas fluorescens with respect to temperature was studied, yielding an Arrhenius plot with a drastic change in slope at 17°C (Fig. 21). Over the cold domain (0–17°C), the temperature characteristic was twofold higher than over the suboptimal domain (17–30°C; Guillou and Guespin-Michel, 1996). The protein content was also measured over the entire temperature range and the authors suggest that, below 17°C, protein degradation is inhibited. This influence of low temperature on protein turnover has also been reported for a psychrotolerant Arthobacter globiformis (Potier, 1990) and could be an explanation for the higher temperature characteristic of the Arrhenius plot in the low-temperature range. A biphasic behavior of the growth rate Arrhenius plot was also reported for a Pseudomonas putida strain (Chablain et al., 1997). It is, however, too early to suggest that this may be a general feature of cold-adapted strains.

239

−2

Ln k = 35.8 − 10691. 1/T

−3

Ln k = 17.5 − 5407. 1/T −4 320

330

340

350

360

370

1/T.10-5 (K-1) Fig. 21. Biphasic Arrhenius plot of the growth rate of Pseudomonas fluorescens. From Guillou and GuespinMichel (1996).

Table 8. Selected doubling times of cold adapted bacteria. Species

TT

Temp. (°C)

dt

Reference(s)

Psychrobacter sp. Str. 1 Frigoribacterium aff. Faeni Rhodococcus sp. Bacillus psychrophilus Bacillus sp. Pseudomonas fluorescens Methanogenium frigidum Yersinia enterocolitica Carnobacterium funditum Vibrio marinus Leuconostoc mesenteroides Leuconostoc citreum Bacillus sp. Psychromonas antarcticus Rhodoferax antarcticus Pseudomonas sp. Clostridium gasigenes Bacillus weihenstephanensis Clostridium algidixylanolyticum Desulfotalea psychrophila Desulfofrigus fragile Enterococcus faecalis Bacillus cereus Methanogenium frigidum Yersinia enterocolitica

Pt Pt Pt Pp Pp Pt Pp Pt Pt Pp Pt Pt Nd Pp Pp Pt Pp Pt Pt Pp Pp Mp Mp Pp Pt

−10 −10 −10 −5 −2 0 0 0 1 3 4 4 5 5 5 10 10 10 10 10 10 10 10 15 15

39d 294d 370d 7h 48h 28h 42d 27h 19h 4h 24h 52h 8h 36h 60h 3h 9h 11h 20h 27h 169h 50h 90h 5d 4h

Bakermans et al., 2003 Bakermans et al., 2003 Bakermans et al., 2003 Morita, 1975 Inniss, 1975 Guillou and Guespin-Michel, 1996 Franzmann et al., 1997 Neuhaus, 2000a Franzmann et al., 1991 Morita and Albright, 1965 Hamasaki et al., 2003 Hamasaki et al., 2003 Brenchley, 1996 Mountfort et al., 1998 Madigan et al., 2000 Morita, 1975 Broda et al., 2000 Prüß et al., 1999 Broda et al., 2000 Knoblauch et al., 1999b Knoblauch et al., 1999b Thammavongs et al., 1996 Prüß et al., 1999 Franzmann et al., 1997 Neuhaus, 2000a

Abbreviations: TT, thermal type; Pp, psychrophilic; Pt, psychrotolerant; Mp, mesophilic; Nd, not determined; and dt, doubling time.

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CHAPTER 1.8 0.05 specific growth rate (h−1)

A

specific growth rate (h−1)

0.006 0.005 0.004

B

0.003 0.002 0.001

0.04 0.03 0.02 0.01 0.00

0.000 −5

0

5

−5

10 15 20 25 30 35 40 temperature (°C)

0

5

10 15 20 25 30 35 40 temperature (°C)

0.25 0.8

(a)

2

0.2

D

0.4

m (h−1)

C

1/td (h−1)

0.6

0.2 0.0

3

0.15 1 0.1 0.05

0

10 20 Temperature (°C)

4

30 0 0

5

10

15

20 25 Temperature,°Ñ

30

35

40

Fig. 22. Thermodependence of growth rates in psychrophilic and psychrotolerant species of bacteria. A) Desulfofaba sp. is an extreme psychrophile. B) Desulfofrigus sp. is a moderate psychrophile. C) Clostridium algidixylanolyticum is a typical psychrotolerant. D) Different species of Acetobacterium are compared. 1) A. bakii (psychrotolerant, from pond sediments); 2) A. paludosum (psychrotolerant, from fen); 3) A. fimetarium (mesophile, from manure); and 4) A. tundrae (psychrophile, from tundra). From Broda et al. (2000), Knoblauch and Jørgensen (1999a), and Nozhevnikova et al. (2001b).

this and other literature. This list gives a rough idea only, and is not a systematic survey. Three plots of growth rate versus temperature are shown in Fig. 22 for psychrotolerant and psychrophilic strains. The kind of mathematical equation that will describe these functions (and why this would be the case) has been discussed widely, but no firm conclusion could be reached (for a review, see, e.g., Berry and Foegeding [1997] and Gounot [1991]). Please note the very different ranges of growth rates shown in Fig. 22. Note also that the terms psychrophilic and psychrotolerant are defined by the growth ranges and by no means reflect growth rates. For instance, a psychrotolerant Pseudomonas has a doubling time of 3 h at 10°C, while a psychrophilic Desulfotalea species grows with a doubling time of 27 h at the same temperature. The growth rate depends on the substrate used in the experiment, among other factors. More important, some bacteria such as Methanogenium or Desulfofrigus are notoriously slow-growing organisms, irrespective of the growth temperature.

Cell Wall A possibly overlooked phenomenon in response to low temperature might be an increase in cell wall thickness. This has been reported for a cold resistant Pseudomonas fluorescens, which showed a 2-fold increase in cell wall thickness, compared to its parent strain, not cold adapted

(Khan et al., 2003). Our own observations during RNA extraction from cold shocked and non-cold shocked Bacillus strains also pointed in such a direction. Cold shocked cells give significantly reduced yields of RNA, with the same protocol. However, similar yields are obtained by elongated bead beating or sonification (pers. observations).

Environment and Applied Aspects The rising interest in cold-adapted microorganisms is fueled by a diverse range of aims connected to their explorations. The answers to the following questions will increase understanding of geo-microbiological processes: Which organisms are found in which environment? And, how does low temperature (Stougaard et al., 2002) and climate (Bidle et al., 2002) influence microbial communities? Other questions deal with treatment of contaminated soil or water, usage of cold-adapted enzymes in technical applications (reviewed by Cavicchioli et al., 2002b), influence of temperature on pathogens (often in relation to food), and finally, usage of cold induced promoters for protein production. A general remark about the occurrence of microbes in the environment was made by Martinus Beijerinck (1851–1931): “Everything is everywhere; the environment selects.” This statement applies to microbial thermotypes.

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Wherever a cold environment is found, e.g., a cold spring (Rudolph et al., 2001), some alpine meadows (Von Stetten et al., 1999), or a refrigerator somewhere in a jungle village, one can be sure that psychroactive microorganisms can be isolated (M. Neuhaus, personal communication). Conversely, hyperthermophilic organisms have been found in soils worldwide, including permafrost regions (Marchant et al., 2002), and some researchers would extend the search area for extremophile microorganisms into extraterrestrial space (Cavicchioli, 2002a; Mitrofanov et al., 2003). The following sections summarize or mention only recent publications in connection with the aims stated above, and one should be aware that each field overlaps.

Environmental Aspects Investigating the ecology of bacteria and archaea is vital to understanding the functioning of the global biochemical cycles. Sulfate reducing bacteria and methanogenic archaea are important terminal oxidizers in the anaerobic mineralization of organic matter and can be seen as ecological equivalents, mineralizing organic matter to CO2 or to CO2 and CH4 in, respectively, highand low-sulfate environments (Purdy et al., 2003). Methanogenesis is also important as a possible climate influence. Methane has a high “green house” effect on the atmosphere. This may explain the great interest in the methanogenesis that occurs in low–temperature environments, which include the sea, the permafrost regions, and deep lakes (Simankova et al., 2003). In low temperature sediments in the Antarctic, Desulfotalea-Desulforhopalus fulforhopalus versus Methanosaeta appear to be the most abundant species of those groups (Purdy et al., 2003). Ecophysiological processes may change in anaerobic systems under extreme conditions (e.g., freezing). In low-sulfate sediments, H2driven methanogenesis was found to be mediated by sulfate reduction. After freezing, both methanogenesis and sulfate reduction decreased. In high-sulfate sediments, sulfate reduction was a major process in frozen and unfrozen samples (Mountfort et al., 2003). In deep lake sediments, a community of psychrophilic methanogens was found, with maximal rates of methane production occurring at 6°C (Nozhevnikova et al., 2003). However, permafrost sediments and other cold environments could also be a sink for methane, since methanotrophic (methane consuming) bacteria have been found in permafrost sediments of Siberia (Khmelenina et al., 2002) and elsewhere (for a review, see Trotsenko and Khmelenina, 2002). In the same environment, anabiotic (dormant) cyst-like bacteria were

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found in sediment samples drawn from a depth of 50–80 m (Dmitriev et al., 2001), and a variety of psychroactive bacteria have been found at a depth of 11–24 m. Interestingly, few Gramnegative isolates could grow at –10°C (the permanent temperature of the sampling site), but all of these isolates grew optimally at around 25– 30°C. Therefore, they have to be classified as psychrotolerant (Bakermans et al., 2003). Removal of soil contamination is an important issue, especially at low temperature. Petroleum hydrocarbons are the most widespread contaminants in the environment. Cold adapted bacteria able to biodegrade such hydrocarbons are already present in pristine soils but increase as a result of the contamination (Margesin et al., 2003). Similarly, psychrotrophic bacteria from the genera Shewanella and Arthrobacter have been isolated from oil-reservoir water and have potential for use in bioremediation (Kato et al., 2001). Obviously, microorganisms exist which are able to degrade the hydrocarbons and other organic wastes under such conditions (Männistö et al., 2001b; Männistö et al., 2001c; Baraniecki et al., 2002; Eriksson et al., 2001; Eriksson et al., 2003; Soares et al., 2003; Thomassin-Lacroix et al., 2001; Thomassin-Lacroix et al., 2002). The poles of the earth (including habitats like sea ice, deep lakes, and similar places) have come into focus. Many scientists are excited by the finding of the big reservoir of liquid water beneath the Antarctic ice shield known as Lake Vostok (Fig. 23). To date, the biological resource of this lake remains untapped, since procedures to remove samples without introducing contamination are still under discussion (Gavaghan, 2002). However, other habitats of Antarctica were examined, and in a permanently frozen

Fig. 23. Lake Vostok (ringed) has lain undisturbed below the ice sheets of Antarctica for many years. From Gavaghan (2002).

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Vostok Station

Ice sheet Borehole

Lake Vostok

Frozen lake water

Bedrock

Sediments

Fig. 24. Schematic view of the borehole to Lake Vostok. It extends beneath Vostok station into areas of frozen lake water but does not reach the lake. From Gavaghan (2002).

Antarctic Lake, a diverse range of phototrophic purple bacteria was found (Karr et al., 2003). This finding is surprising because organisms in Antarctic habitats commonly face continuous low temperatures, as well as poor light conditions and nutrient limitations, making Antarctica appear deserted. But many adapted organisms are thriving below the snow, playing a most fundamental role in the polar ecosystem (for reviews, see Laybourn-Parry [2002]; Thomas and Dieckmann [2002b], and Rossi et al. [2003]).

Technical Uses An important step in wastewater treatments is the removal of water pollutants by microorganisms. But even in a moderate climate, wastewater temperature may drop to 10°C or 15°C in winter, eventually inhibiting growth of the microbial flora. Under certain conditions low temperature might be beneficial (e.g., low temperature reduces the number of bacteria introduced into the sea by Antarctic research stations; Hughes and Blenkharn, 2003), but mainly it poses a challenge to modern wastewater treatment facilities. Different technical

solutions have been proposed to treat wastewater successfully at 13°C (for a review, see Lettinga et al., 2001). Another problem is the huge amount of solids entering sewage treatment facilities. Anaerobic digestion might decrease the amount, but this leads to fouling and biogas (including significant amounts of methane) emission (Nozhevnikova et al., 2001a). Conversely, psychrotolerant nitrifying bacteria may pose a threat to drinking water quality in cold climates (Lipponen et al., 2002). For extraction of artificially expressed proteins, either for the laboratory (Moran et al., 2001) or for technical uses (Tutino et al., 2001), cold adapted organisms or promoters activated at low temperatures might have certain advantages for the production of thermolabile, toxic, or proteolytically sensitive proteins, for increasing proper folding, increased solubility, or stability (Gonzalez et al., 2003; Mujacic et al., 1999; Takeuchi et al., 2003; Tutino et al., 2001). For more information about the usage of cold inducible promoters in E. coli, the reader is referred to the review by Baneyx (1999) or the methodological papers by Baneyx and Mujacic (2003), Qing et al. (2004), and Duilio et al. (2004).

Food Production and Protection Psychrophilic and psychrotolerant microorganisms are of great importance to the food industry. These organisms are used for direct production, e.g., of dairy products, on the one hand, and may spoil cold stored food or be pathogenic on the other (Russell, 2002). Lactic acid bacteria (LAB) are traditionally used to produce fermented food, but this heterogeneous group is also used for other purposes, e.g., as probiotics and bioprotectives. Because of the importance and amount of food processed with LAB, a huge body of literature has accumulated that is focused on the cold shock response and cold survival. Different methods are applied to enhance survival of LAB after cold shock or freezing of starter cultures. Exopolysaccharides, overproduction of MCSPs, and employment of other stressful conditions enhances survival of LAB after chilling or freezing (Derzelle et al., 2003; Hong and Marshall, 2001; Maus and Ingham, 2003; Serror et al., 2003a). The discovery of new thermosensitive replicons and two transposons by Serror et al. (2003b) added to the toolbox for manipulating Lactobacillus species. For a review of the stress responses in LAB, including cold shock, see Van de Guchte et al. (2002) and the extensive literature survey provided by Carr et al. (2002). Several studies have been published on the occurrence of psychrotrophic and psychrophilic

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bacteria in food matrices. The publications can be divided into those dealing with spoiling organisms and those dealing with survival of certain pathogens under different low temperature conditions. A few recent publications in this field of low temperature research will be briefly mentioned. The smoky odor in chocolate milk stored at 4–9°C is due to guaicol produced by the spoiling organism Rahnella aquatilis. This is the first identification of an organism responsible for this type of spoilage (Jensen et al., 2001). Psychrotolerant LAB have been identified as causative agents for spoilage in cooked meat products (Hamasaki et al., 2003). The main flora of cold stored pork meat was found to be Pseudomonas, Aeromonas and Acinetobacter species (Olsson et al., 2003). Neither cold nor carbon dioxide induce a viable but nonculturable state in Listeria monocytogenes (Li et al., 2003). However, reduction in the number of this organism is achieved by using essential oils and freezing (Cressy et al., 2003). Bacillus cereus can be controlled in chilled dairy products by adding variacin, a lantibiotic produced by Kocuria varians (O’Mahony et al., 2001). However, the resistance of Salmonella and E. coli OH157:H7 (EHEC) to other stresses was increased by subjecting cells to cold temperature beforehand (Bollman et al., 2001; Gawande and Bhagwat, 2002). This increased resistance can be used to advantage in that more viable counts can be obtained by classical plating methods, thereby increasing the sensitivity of pathogen detection (Sol et al., 2002). Other organisms of particular concern able to grow at low temperature include Clostridium perfringens, Campylobacter jejuni, Staphylococcus aureus, Yersinia enterocolitica, etc. (Chan et al., 2001; Guentert and Linton, 2003; Harrison et al., 2000; Kalinowski et al., 2003; Steele and Wright, 2001; Zhao et al., 2003). Interestingly, the source of psychrophilic clostridia spoiling vacuum packed chilled meat products is most likely soil particles and fecal material introduced at the abattoir (Boerema et al., 2003). Reduction or elimination of pathogenic psychrotolerant bacteria is a major aim of food processing. However, the ecology of the pathogen growing in the food matrix is often poorly understood. An elegant in situ method for monitoring a pathogenic Yersinia enterocolitica in cheese samples was reported by (Maoz et al., 2002). A full-length luxCDABE operon was introduced in the genome of this organism, which carried a constitutive promoter. The emitted light, corresponding to colony forming unit (cfu) counts, was monitored with a sensitive, charge coupled device (CCD) camera. This system does not need the addition of any further substance like antibiotics (to maintain a plasmid) or substrate for the light producing LuxAB enzymes. The influence

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of, e.g., bioprotective cultures and other means to control the pathogen, can be monitored in situ without laborious cfu plate countings. This technique can be used to monitor other pathogenic organisms (Francis et al., 2000).

Pathogens Low temperature in connection with pathogens again includes many different facets. Survival of psychrotolerant pathogenic organisms in food has been mentioned in the section above. In this section, some recent findings about the connection between low temperature and pathogens and virulence are highlighted. Temperature regulation of virulence factors has been reviewed by DiRita et al. (2000) and Konkel and Tilly (2000). Many pathogens regulate virulence genes via temperature sensing mechanisms (see the section Bacterial Cold Sensors in this Chapter). Yersinia enterocolitica contains a virulence plasmid, which carries multiple regions of intrinsic curvature. These bends are detectable at 30°C but melt at 37°C, the temperature at which the cells undergo phenotypic switching (Rohde et al., 1999). Other examples of such a behavior can be found in Vibrio salmonicida. Disease of Atlantic salmon occurs only if the water temperature is below 10°C. An important virulence factor of this species might be iron siderophores and other iron uptake systems expressed only at the low temperature (Colquhoun and Sorum, 2001). E.g. the main cold shock protein in L. monocytogenes seems to be a ferritin-like protein. A similar finding has been reported from Streptococcus thermophilus, also expressing an iron binding protein after cold shock (Hébraud, 2000 #2662; Nicodeme, 2004 #2525). In L. monocytogenes, the transcriptional activator PrfA controls many virulence genes. The mRNA of this activator acts as thermometer. The 5′ untranslated region renders the ribosome binding site inaccessible at lower temperature (e.g., 30°C) and then switches to an accessible form at higher temperature (e.g., 37°C; Johansson et al., 2002). However, findings showed virulence gene expression at 30°C and below in artificially infected Drosophila melagonaster (Mansfield et al., 2003). Phytopathogens (e.g., Pseudomonas syringae) were also found to regulate virulence factors in response to temperature. The phytotoxin coronatine mimics the plant hormone jasmonate. The biosynthesis cluster is regulated by a twocomponent system, probably sensing membrane fluidity (Smirnova et al., 2002). Thermoregulated expression of virulence factors in plantassociated bacteria has been reviewed by Smirnova et al. (2001a). Conversely, a distinct

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Table 9. Recent reviews in connection with life at low temperatures in bacteria Scope Enzymes Enzymes Enzymes Enzymes Enzymes Enzymes Food Food Food Food General General General General Geology Geology Geology Geology Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Molecular Pathogens Pathogens Technique Technique Technique

Title Molecular basis of cold adaptation Molecular adaptations to cold in psychrophilic enzymes Psychrophilic enzymes: hot topics in cold adaptation Cold-adapted enzymes: from fundamentals to biotechnology Psychrophilic enzymes: revisiting the thermodynamic parameters of activation may explain local flexibility Toward a molecular understanding of cold activity of enzymes from psychrophiles A review of aerobic and psychrotrophic plate count procedures for fresh meat and poultry products Bacterial membranes: the effects of chill storage and food processing. An overview Stress responses in lactic acid bacteria The role of cold-shock proteins in low-temperature adaptation of food-related bacteria Cold shock response and adaptation at near-freezing temperature in microorganisms Biology of extremophilic and extremotolerant methanotrophs Life at low temperature Extremophiles 2002 Extremophiles and the search for extraterrestrial life Life in the deep freeze Survival mechanisms in Antarctic lakes Antarctic Sea ice—a habitat for extremophiles Temperature sensing and cold acclimation The link between bacterial radiation resistance and cold adaptation Environmental sensing mechanisms in Bordetella Low-temperature sensors in bacteria Bacterial cold shock proteins Bacterial cold-shock response at the level of DNA transcription, translation and chromosome dynamics Transcriptional and post-transcriptional control of cold-shock genes Conservation of the cold shock domain protein family in plants Identifying global regulators in transcriptional regulatory networks in bacteria Recent developments in bacteria cold-shock response Regulation of the desaturation of fatty acids and its role in tolerance to cold and salt stress Biliproteins and phycobilisomes from cyanobacteria and red algae at the extremes of habitat Control of transcription termination in bacteria by RNAbinding proteins that modulate RNA structures Cold adaptation of archaeal elongation factor 2 (EF-2) proteins. Coping with the cold: the cold shock response in the Grampositive soil bacterium Bacillus subtilis Bacterial cold shock responses Molecular components of physiological stress responses in Escherichia coli Virulence gene regulation inside and outside Temperature-regulated expression of bacterial virulence genes Cold-inducible promoters for heterologous protein expression Low-temperature extremophiles and their applications Challenge of psychrophilic anaerobic wastewater treatment

Reference(s) D’Amico et al., 2002a Feller, 2003a Feller and Gerday, 2003b Gerday et al., 2000 Lonhienne et al., 2000 Russell, 2000 Jay, 2002 Russell, 2002 Van de Guchte et al., 2002 Wouters et al., 2000b Inouye and Phadtare, 2004 Trotsenko and Khmelenina, 2002 Neuhaus and Scherer, 2004 Rossi et al., 2003 Cavicchioli, 2002a Gavaghan, 2002 Laybourn-Parry, 2002 Thomas and Dieckmann, 2002b Browse and Xin, 2001 Chattopadhyay, 2002 Coote, 2001 Eriksson et al., 2002 Ermolenko and Makhatadze, 2002 Golovlev, 2003 Gualerzi et al., 2003 Karlson and Imai, 2003 Martinez-Antonio and Collado-Vides, 2003 Phadtare, 2004a Sakamoto and Murata, 2002 Samsonoff and MacColl, 2001 Stülke, 2002 Thomas and Cavicchioli, 2002a Weber and Marahiel, 2002 Weber and Marahiel, 2003 Wick and Egli, 2004 DiRita et al., 2000 Konkel and Tilly, 2000 Baneyx and Mujacic, 2003 Cavicchioli et al., 2002b Lettinga et al., 2001

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group of psychrotrophic Pseudomonas species can protect plants by disease-suppression or growth promotion (Johansson and Wright, 2003; Katiyar and Goel, 2003; Mishra and Goel, 1999). One could speculate that this increased plant resistance is due to CspA, which was shown to be a highly active elicitor of tobacco defense responses (Felix and Boller, 2003). Recently, the general involvement of cold shock genes in virulence has been suggested. Cold-inducible RNases (PNPase and RNase H) were found to be important for full virulence of Shigella and enteroinvasive E. coli (see Cairrão et al. [2003] and references therein). The PerR regulon of Streptococcus pyogenes is needed for full virulence and contains a Csp (Brenot et al., 2005). Two studies showed a connection between susceptibility to certain antimicrobial substances and the cold shock proteins in Staphylococcus aureus: Methicillin resistant S. aureus can be treated with the detergent Triton X-100, which reduces the methicillin-resistance. The more resistant a particular strain was before treatment the more its resistance decreased. Comparative proteomics revealed that the MCSPs CspABC of such methicillin-resistant strains (unlike methicillin-sensitive strains) were highly induced (Cordwell et al., 2002). Insertion of a transposon in the cspA gene increases the resistance to an antimicrobial peptide of human cathepsin G in the same bacterium (Katzif et al., 2003). Possibly, increased MCSP levels increase the susceptibility to antibacterial substances in S. aureus, but more evidence is needed to define the role that CspA plays in such substance resistance. This finding is for some reason contrary to the cross-resistance to other various stresses, which normally increases after induction of the cold shock response and vice versa.

Concluding Remarks The large number of proteins synthesized upon cold shock as well as in cold acclimation of psychrotolerant microorganisms (see the references in the sections The Cold Shock Response and Cold Acclimation in this Chapter) is clear evidence that many cellular processes contribute to a bacterium’s capacity for growth at low temperature. Also obvious is that adaptations at the structural level of rRNA and proteins as well as transient adaptations in the pattern of gene expression are involved in cold adaptation. In toto, a variety of processes thus affect the fitness of cold-adapted bacteria, which, in turn, are important to understand the role of these organisms in their habitat. However, our understanding of the molecular structure of cold adaptation is still in its initial stage. Consequently, our

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understanding of the evolution of cold tolerance is also quite sketchy. The frequent presence of cold tolerant and mesophilic strains in the same genus or even in the same species suggests that the evolution of psychrotolerance is most probably a multiple step process, which may have occurred many times in parallel. Even though this review tries to cover many aspects of bacterial life at low temperature, not every area can be covered in detail. Many specialized reviews have been written in recent years. Most of these reviews were cited in the appropriate section above, but to simplify the search for a specific topic, the following table lists reviews or similar articles with a broader scope (Table 9).

Literature Cited Abd El-Rahman, H. A., D. Fritze, C. Sproer, and D. Claus. 2002. Two novel psychrotolerant species, Bacillus psychrotolerans sp. nov. and Bacillus psychrodurans sp. nov., which contain ornithine in their cell walls. Int. J. Syst. Evol. Microbiol. 52:2127–2133. Agafonov, D. E., V. A. Kolb, I. V. Nazimov, and A. S. Spirin. 1999. A protein residing at the subunit interface of the bacterial ribosome. Proc. Natl. Acad. Sci. USA 96:12345–12349. Agafonov, D. E., V. A. Kolb, and A. S. Spirin. 2001. A novel stress-response protein that binds at the ribosomal subunit interface and arrests translation. Cold Spring Harb. Symp. Quant. Biol. 66:509–514. Aguilar, P. S., J. E. Cronan Jr., and D. de Mendoza. 1998. A Bacillus subtilis gene induced by cold shock encodes a membrane phospholipid desaturase. J. Bacteriol. 180: 2194–2200. Aguilar, P. S., P. Lopez, and D. de Mendoza. 1999. Transcriptional control of the low-temperature-inducible des gene, encoding the delta5 desaturase of Bacillus subtilis. J. Bacteriol. 181:7028–7033. Aguilar, P. S., A. M. Hernandez-Arriaga, L. E. Cybulski, A. C. Erazo, and D. de Mendoza. 2001. Molecular basis of thermosensing: A two-component signal transduction thermometer in Bacillus subtilis. EMBO J. 20:1681– 1691. Akila, G., and T. S. Chandra. 2003. A novel cold-tolerant Clostridium strain PXYL1 isolated from a psychrophilic cattle manure digester that secretes thermolabile xylanase and cellulase. FEMS Microbiol. Lett. 219:63–67. Albers, S. V., J. L. van de Vossenberg, A. J. Driessen, and W. N. Konings. 2000. Adaptations of the archaeal cell membrane to heat stress. Front. Biosci. 5:D813–D820. Allen, E. E., D. Facciotti, and D. H. Bartlett. 1999. Monounsaturated but not polyunsaturated fatty acids are required for growth of the deep-sea bacterium Photobacterium profundum SS9 at high pressure and low temperature. Appl. Environ. Microbiol. 65:1710–1720. Allen, E. E., and D. H. Bartlett. 2000. FabF is required for piezoregulation of cis-vaccenic acid levels and piezophilic growth of the deep-sea bacterium Photobacterium profundum strain SS9. J. Bacteriol. 182:1264–1271. Allen, E. E., and D. H. Bartlett. 2002. Structure and regulation of the omega-3 polyunsaturated fatty acid synthase

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Prokaryotes (2006) 2:263–282 DOI: 10.1007/0-387-30742-7_9

CHAPTER 1.9 e f iL

ta

hg iH

t l aS

sno i t ar tnecnoC

Life at High Salt Concentrations AHARON OREN

Introduction A great variety of prokaryotes, Bacteria as well as Archaea, can be found in saline and hypersaline environments. These microorganisms are adapted to life at high salt concentrations and to the high osmotic pressure of their environment resulting from the high salinity. This chapter presents a general overview of the hypersaline environments as biotopes for prokaryotic life, the types of organisms encountered in them, and the mechanisms the different groups of prokaryotes have developed to cope with the special requirements of life in the presence of molar concentrations of salt. More detailed information on the variety of halophilic organisms can be found in the specific chapters that deal with the different taxonomic groups.

Saline and Hypersaline Habitats The greatest part of the biosphere is saline. The waters of the oceans and seas that cover most of the earth’s surface contain around 35 g dissolved salts per liter. Higher salt concentrations are often encountered in near-shore environments such as salt marshes, sabkhas and lagoons, under conditions in which evaporation is rapid and water exchange with the open sea is slow. Still higher concentrations of salts, up to saturation of NaCl and beyond, exist in natural inland salt lakes such as the Dead Sea on the border between Israel and Jordan (with total dissolved salts of about 340 g/liter), the Great Salt Lake in Utah (up to 333 g/liter in 1975, although values have decreased since 1975 due to the positive water balance of the lake), and many others. Gradients of increasing salt concentrations are found in the man-made evaporation ponds and crystallizer basins of multi-pond solar saltern systems near tropical and subtropical shores worldwide. All these environments, from seawater salinity to NaCl-saturated brines, are potential habitats for prokaryotic life (Brock, 1979; Rodriguez-Valera, 1988, 1993). Additional hypersaline environments inhabited by salt-tolerating (halotolerant)

and salt-loving (halophilic) microorganisms are salted food products such as salted fish, animal hides treated with salt for their preservation, saline soils, and subterranean brines that are often associated with oil fields. The properties of hypersaline environments as habitats for halophilic and halotolerant prokaryotes are primarily defined according to the total salt concentration. However, also the ionic composition is a key factor determining the properties of the environment as a biotope. Brines that originated by evaporation of seawater (so-called thalassohaline brines) reflect the ionic composition of the sea, at least during the first stages of evaporation (Fig. 1). The ionic composition starts to change significantly when evaporation proceeds to the stage at which the solubility limit of CaSO4 is reached and gypsum precipitates (at a total salt concentration above 100–120 g/liter). The brines that enter saltern crystallizer ponds in multi-pond salterns are thus depleted in calcium and to a minor extent in sulfate. During the subsequent precipitation of NaCl as halite, the ionic composition changes again, and the relative concentrations of K+ and Mg2+ increase. The Great Salt Lake, Utah, though since long detached from the world ocean, still reflects in its ionic composition the seawater that contributed its salt, and therefore its waters can still be classified as thalassohaline. Thalassohaline brines are characterized by neutral or slightly alkaline pH values (7–8). In other hypersaline environments, the ionic composition may greatly differ from that of seawater (“athalassohaline environments”). The Dead Sea is a prime example of an athalassohaline lake. Here divalent cations dominate, with concentrations of Mg2+ (1.89 M) and Ca2+ (0.45 M) exceeding those of Na+ (1.56 M) and K+ (0.20 M) (1998 values). As a result of the high Ca2+ concentration the solubility of sulfate is low, and monovalent anions (Cl- and Br-) make up more than 99.9% of the anion sum (Fig. 1). The pH of the Dead Sea brine is relatively low, around 5.8–6.0. Alkaline athalassohaline brines are relatively abundant. Alkaline “soda lakes” are present in

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CHAPTER 1.9

Concentration (mol% of total cations or anions)

Seawater 100

NaCl-Saturated Great Salt Lake Evaporated Seawater CL–

CL– Na+

Dead Sea CL–

CL–

Lake Magadi Na+

Wadi Natrun Lake Zugm Na+

Na+

Na+

CO32–

80

CL–

60 Mg2– Na+

40

CL–

Na+

CO32–

20 Mg2+

K+ Ca2+

SO42–

Mg2+ K+ Ca2+

SO42–

Mg2+ + K Ca2+

Ca2+ SO42–

K+ Br

K+

K+

SO42–

0 Fig. 1. The ionic composition of seawater and selected hypersaline environments. The bars represent the mol fraction of different cations and anions in the brines. Data for seawater, saltern brines and Great Salt Lake were derived from Javor (1989), and data on Lake Magadi (analyses for 1976) and on Lake Zugm, Wadi Natrun, Egypt, were from Grant and Tindall (1986) and from Grant et al. (1998a), respectively. Dead Sea data (deep water, 1998) were obtained from Michael Beyth (the Israel Ministry of National Infrastructures, personal communication).

diverse geographic locations such as in East Africa (Lake Magadi and other lakes in Kenya and Tanganyika), in the Wadi Natrun in Egypt, and in California, Nevada, India, Tibet, China, and elsewhere. Here the salt composition is dominated by monovalent cations. Because of the high pH (up to 10–11 and higher) the solubility of the divalent cations Mg2+ and Ca2+ is very low, and the concentrations of these ions may be below the detection limit. Carbonate and bicarbonate ions contribute a significant part of the anion sum in such lakes, in addition to chloride and sulfate.

Classification and Phylogeny of Prokaryotes Living at High Salt Concentrations Microorganisms adapted to life at high salt concentrations are widespread, both within the bacterial and the archaeal domain. As a result, highly diverse prokaryote communities can be found at all salt concentrations from seawater up to about 340–350 g/liter (brines saturated with NaCl) in both thalassohaline and athalassohaline environments. A few microorganisms can adapt to life over the whole salt concentration range from near fresh water to halite saturation. Halomonas elongata is a well-known example of such a bacterium (Vreeland et al., 1980). In most cases, however, each organism has a relatively restricted salt concentration range enabling growth. Some bacteria are adapted to life in sat-

urated and near-saturated brines, being unable to grow and even survive at NaCl concentrations below 15–20%. Most representatives of the halophilic Archaea of the order Halobacteriales show such a behavior. Others thrive at an intermediate salt concentration range. Salt requirement and tolerance may be temperature-dependent, and many cases have been described in which both salt tolerance and requirement are enhanced at increased temperatures (see e.g., Mullakhanbhai and Larsen, 1975). Different classification schemes have been designed to define the salt relationships of microorganisms. All such schemes are artificial to some extent. Because of the continuum of properties found within the prokaryote world there will always be organisms that cannot unequivocally be classified within any of the groups defined. The most widely accepted classification according to salt dependence and salt tolerance is that of Kushner (1978, 1985), given in a slightly modified form in Table 1. This scheme recognizes different degrees of salt dependence (slightly, moderately, and extremely halophilic). In addition, halotolerant microorganisms exist that, while not requiring high salt concentrations for growth, are able to grow at high concentrations of NaCl and of other salts. Staphylococcus species present a good example for this category, as they grow well both in the absence of salt and at NaCl concentrations as high as 10–15% and even higher, a property often exploited in the design of selective and diagnostic growth media. It should be noted that classification should be based not only on

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Table 1. Classification of microorganisms according to their response to salt. Properties

Examples

Grows best in media containing less than 0.2M salt Grows best in media containing 0.2–0.5M salt Grows best in media containing 0.5–2.5M salt Grows best in media containing 1.5–4.0M salt Grows best in media containing 2.5–5.2M salt Non-halophile which can tolerate salt; if the growth range extends above 2.5M salt, it may be considered extremely halotolerant

Most freshwater bacteria Most marine bacteria Salinivibrio costicola Halorhodospira halophila Halobacterium salinarum Staphylococcus aureus

Some of the early studies on this unique group of prokaryotes were summarized in Larsen’s classic essay on “the halobacteria’s confusion to biology” (Larsen, 1973), and a full account of their properties can be found elsewhere (e.g., Kushner, 1985; Oren, 1994; Tindall and Trüper, 1986). The presence of dense communities of members of the Halobacteriales in hypersaline environments often can be observed with the unaided eye thanks to the bright red, orange, or purple coloration of most representatives of the group and to the extremely high community densities at which these Archaea may develop. The occurrence of red hues has been documented for the north arm of the Great Salt Lake (Post, 1977), the Dead Sea (Oren, 1988a), and hypersaline alkaline lakes such as Lake Magadi, Kenya (Grant and Tindall, 1986). Red colored brines also are present typically during the final stages of the evaporation of seawater in solar saltern crystallizer ponds (Borowitzka, 1981; Javor, 1989; Oren, 1993, 1994) (Figs. 3–6). Sometimes other types of microorganisms may also contribute to the color of the brine, such as the bcarotene-rich, green halophilic alga Dunaliella salina in saltern ponds (Fig. 6), or photosynthetic purple bacteria of the genus Ectothiorhodospira

m

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Clostridium Bacillus cterium Helioba a acteri ive B -Posit

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Fig. 2. Phylogenetic tree of the Bacteria and the Archaea, based on 16S rRNA sequence comparisons, indicating the distribution of halophilism. Bold lines indicate branches containing representatives able to grow at or near optimal rates at NaCl concentrations exceeding 15%.

cte r te ba ac lavo F e Fl xib

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BACTERIA

occus Thermoc Methanococcus jannaschii

the behavior toward NaCl but to other ions as well, especially for organisms adapted to life in athalassohaline environments (Edgerton and Brimblecombe, 1981). The table is based on classification schemes proposed by Kushner (1978, 1985). Halophilic behavior is found all over the phylogenetic tree of the prokaryotes, both within the Archaea and the Bacteria. Within the archaeal domain, growth at salt concentrations above 15–20% has been documented not only in the Halobacteriales but also in the methanogenic genus Methanohalophilus (family Methanosarcinaceae). Most halophilic Bacteria characterized belong to the g-subdivision of the Proteobacteria, but moderate halophiles can also be found in other subgroups of the Proteobacteria, the low G+C and the high G+C Gram-positive Bacteria, the cyanobacterial branch, the Flavobacterium branch, and the Spirochetes (Fig. 2) (Ventosa et al., 1998). The archaeal order of the Halobacteriales contains the extreme halophiles par excellence. These are highly specialized microorganisms, most of which will not grow at total salt concentrations below 2.5–3 M. When suspended in solutions containing less than 1–2 M salt, cells are irreversibly damaged, and many species will lyse.

γ-Pro teoba cteria β- P rote oba cte ria Pl an cto my ce s

Category Non-halophilic Slight halophile Moderate halophile Borderline extreme halophile Extreme halophile Halotolerant

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Fig. 3. Saltern crystallizer pond of the Israel Salt Company at the Red Sea coast near Eilat at a total dissolved salt concentration of about 340 g/liter, colored red by halophilic Archaea.

or Halorhodospira, which may contribute at least part of the red coloration of the brines in the alkaline lakes of the Wadi Natrun, Egypt (Jannasch, 1957). Halophilic Archaea may survive for prolonged periods within halite crystals. This property has aroused considerable interest in recent years, following the isolation of viable halophilic Archaea from salt collected from salt mines dating from the Triassic (195–225 million years B.P.) and Permian (225–270 million years B.P.) periods (Norton et al., 1993). A variety of halophilic Archaea was recently isolated from the Permian Salado salt formation near Carlsbad, New Mexico, including many unknown types (Vreeland et al., 1998). Controversy still exists over whether these bacteria were trapped within

Fig. 4. Saltern crystallizer pond of the Israel Salt Company at the Mediterranean coast near Atlit at a total dissolved salt concentration of about 340 g/liter, colored red by halophilic Archaea.

CHAPTER 1.9

Fig. 5. Saltern ponds of the Cargill Solar Salt Works (Newark, CA), showing a crystallizer pond colored brightly red by halophilic Archaea (courtesy of Carol D. Litchfield, George Mason University, Fairfax, VA).

the crystals, where they retained their viability, or whether these cells may have entered the salt more recently, during disturbances of the salt layer that were caused by natural phenomena or human activity. Vreeland and Powers (1999) give a critical discussion of the intriguing findings of viable prokaryotic cells within ancient salt deposits. Recently, the heterogeneous 16S rRNA genes in Haloarcula isolates from ancient salt deposits have been compared with those of modern strains in the expectation that there are fewer differences between the genes of truly ancient Haloarcula than in modern strains if the gene multiplicity originated by duplication. No indications were found that the genes from present-day strains are more divergent than the ancient ones (Grant et al., 1998b).

Fig. 6. Brines in crystallizer pond of the Cargill Solar Salt Works (Newark, CA), colored in part red due to dense communities of halophilic Archaea (foreground) and in part showing a more orange color imparted by the b-carotene-rich unicellular green alga Dunaliella salina. The total dissolved salt at the time (February 1997, following a period of heavy rains) was about 250 g/liter.

CHAPTER 1.9

Other taxonomically coherent groups consisting solely or mainly of halophilic microorganisms are the order Haloanaerobiales and the family Halomonadaceae. The Haloanaerobiales form an order of moderately halophilic anaerobic bacteria within the low G+C branch of the Gram-positive Bacteria (Rainey et al., 1995). As discussed below and elsewhere (Oren, 1992), this group is of special interest because the mechanism of salt adaptation used by its members resembles that of the aerobic halophilic Archaea rather than that of the other halophilic or halotolerant Bacteria. The family of the Halomonadaceae (g-subgroup of the Proteobacteria) contains some of the most versatile prokaryotes with respect to their adaptability to a wide range of salt concentrations. The adaptations of some of its representatives to salt have been studied extensively (Franzmann et al., 1988b; Ventosa et al., 1998).

Thermophilic, Psychrophilic, and Alkaliphilic Halophiles Among the halophilic prokaryotes some are adapted to other forms of environmental stress in addition to salt stress. Thus, thermophilic, psychrophilic, and alkaliphilic halophiles are known. No acidophilic halophiles have been described as yet. The Dead Sea with a pH of about 6.0 is probably the most acidic environment in which mass development of halophilic Archaea has been reported (Oren, 1988a). Most aerobic halophilic Archaea of the order Halobacteriales have rather high temperature optima, in the range between 35 and 50∞C and sometimes even higher. Growth at high temperatures may be an adaptation to the often relatively high temperatures of salt lakes in tropical areas. Within the anaerobic Bacteria of the order Haloanaerobiales several moderately thermophilic representatives were described. Halothermothrix orenii, the first truly thermophilic halophile discovered, was isolated from Chott El Guettar, a warm saline lake in Tunisia. It grows optimally at 60∞C and up to 68∞C at salt concentrations as high as 200 g/liter (Cayol et al., 1994). Acetohalobium arabaticum strain Z-7492 has a temperature optimum of 55∞C (Kevbrin et al., 1995). Cold-adapted halophiles also occur. The halophilic Archaeon Halorubrum lacusprofundi was isolated from Deep Lake, Antarctica, a hypersaline lake in which the water temperature varies according to the season between below zero to +11.5∞C. The isolate grows optimally at 31–37∞C, but slow growth does occur down to tempera-

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tures of 4∞C (Franzmann et al., 1988a). In addition, a variety of halophilic and halotolerant Bacteria was isolated from different salt lakes in Antarctica (Dobson et al., 1991; Franzmann, 1991; McMeekin and Franzmann, 1988; McMeekin et al., 1993). Halophilic Archaea of the order Halobacteriales are abundant in hypersaline soda lakes such as Lake Magadi (Kenya) (Grant and Tindall, 1986; Tindall and Trüper, 1986; Tindall et al., 1980, 1984), the Wadi Natrun lakes (Egypt) (Imhoff et al., 1978, 1979; Soliman and Trüper, 1982), and soda lakes in China (Wang and Tang, 1989) and India (Upasani and Desai, 1990). They may impart a red color to such lakes. These environments are characterized by salinity at or close to halite saturation and contain, in addition, high concentrations of carbonates. The pH values are around 10–11 (Grant and Tindall, 1986). Also anaerobic halophilic alkaliphiles occur in such environments. Lake Magadi was shown to harbor a varied anaerobic community, including cellulolytic, proteolytic, saccharolytic, and homoacetogenic bacteria (Shiba and Horikoshi, 1988; Zhilina and Zavarzin, 1994; Zhilina et al., 1996). The homoacetogen Natroniella acetigena was isolated from this environment. Its pH optimum is 9.8–10.0, and it can grow up to pH 10.7 (Zhilina et al., 1996). Anaerobes were also isolated from the alkaline saline Big Soda Lake, Nevada (Shiba and Horikoshi, 1988; Shiba et al., 1989).

Metabolic Diversity of Halophilic Microorganisms A survey of the halophilic microorganisms for metabolic diversity shows that many, but not all types of dissimilatory metabolism known within the prokaryotic world, can function in hypersaline environments as well. Figure 7 presents an overview of the functional diversity of halophilic prokaryotes, based both on laboratory experiments with isolated cultures and on measurements of the processes as they occur in nature. Oxygenic photosynthesis by cyanobacteria can occur almost up to NaCl saturation. Though the main planktonic primary producers in most hypersaline environments are eukaryotic algae of the genus Dunaliella (Javor, 1989; Oren, 1988a, 1994; Post, 1977), cyanobacteria such as Aphanothece halophytica [Cyanothece; for a discussion of the problems in the taxonomy of the “Halothece” group see Garcia-Pichel et al. (1998)] are often found abundantly in benthic microbial mats that cover the shallow sediments of salt lakes and saltern ponds, especially in the salinity range between 150–250 g/liter (Oren,

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CHAPTER 1.9 Salt concentration (g/l) 0

100

Oxygenic photosynthesis Anoxygenic photosynthesis Aerobic respiration Denitrification Fermentation Proton-reducing acetogens

?

Dissimilatory sulfate reduction - incomplete oxidizers Dissimilatory sulfate reduction - complete oxidizers Methanogenesis from H2 + CO2 Methanogenesis from acetate

?

Methanogenesis from methylated amines Acetate formation from H2 + CO2

200

300

Fig. 7. Approximate upper salt concentration limits for the occurrence of selected microbial processes. Values presented are based in part on laboratory studies of pure cultures (black bars) and on activity measurements of natural microbial communities in hypersaline environments (white bars). Data were derived in part from Brandt and Ingvorsen (1998) , Ollivier et al. (1998), Oremland and King (1989), Oren (1988b, 1999), Sokolov and Trotsenko (1995), Zhilina and Zavarzin (1990), and many additional sources.

Chemolithotrophic oxidation of sulfur compounds Autotrophic ammonia oxidation Autotrophic nitrite oxidation

?

Aerobic methane oxidation

Fig. 8. A crust of gypsum densely populated with cyanobacteria (Aphanothece halophytica [Cyanothece; see GarciaPichel et al., 1998] and others) in a saltern evaporation pond of the Israel Salt Company at the Red Sea coast near Eilat at a total dissolved salt concentration of 286 g/liter. The author is sampling the carotenoid-rich upper unicellular cyanobacterial layer and the green layer of filamentous and unicellular cyanobacteria below.

1999). Figure 8 shows an example of a dense benthic community of unicellular cyanobacteria living within a gypsum crust of a saltern pond at a salt concentration of 286 g/liter. Anoxygenic photosynthetic prokaryotes also abound up to the highest salt concentrations. Examples are representatives of the genus Halorhodospira, which contains species such as Halorhodospira halochloris and Halorhodospira halophila that can be classified as borderline extreme halophiles (see Table 1). Halorhodospira cells were documented to impart a bright red color to the alkaline hypersaline lakes of Wadi Natrun, Egypt (Jannasch, 1957). Addi-

tional halophilic purple bacteria have been characterized, such as Halochromatium and Thiohalocapsa (Caumette et al., 1991, 1997; Imhoff et al., 1998). There is also a report that green sulfur bacteria (Chlorobium or a relative) may occur at high salt concentrations (Anderson, 1958), but details are lacking. Most halophilic prokaryotes that have been isolated and studied are aerobic chemoorganotrophs. Aerobic breakdown of organic compounds is possible at salinities up to NaCl saturation. Both halophilic Archaea of the order Halobacteriales and different types of Bacteria may be involved in the breakdown of organic compounds in hypersaline environments. In addition to simple compounds such as sugars, amino acids, etc., also a number of unusual substrates can be degraded at high salt concentrations. Aliphatic and aromatic hydrocarbons, including even- and odd-carbon-number saturated hydrocarbons; saturated isoprenoid alkanes (pristane); different aromatic compounds, including benzoate, cinnamate, and phenylpropionate; and long-chain fatty acids, such as palmitic acid, have been shown to serve as sole carbon and energy sources for certain isolates of halophilic Archaea (Bertrand et al., 1990; Emerson et al., 1994; Kulichevskaya et al., 1991). Degradation of hexadecane was shown in the Great Salt Lake up to a salinity of 172 g/liter (Ward and Brock, 1978). Other unusual compounds shown to be degraded or transformed at high salt concentrations are formaldehyde (Azachi et al., 1995) and organophosphorus compounds (DeFrank and Cheng, 1991). Oren et al. (1992) presented an overview of the potential for breakdown of unusual compounds, including industrial pollutants, at high salt concentrations.

CHAPTER 1.9

Oxygen is poorly soluble in concentrated brines, and therefore it is not surprising to find a considerable variety of anaerobic halophilic heterotrophs. Many representatives of the aerobic halophilic Archaea of the order Halobacteriales can grow anaerobically by using nitrate as electron acceptor (Mancinelli and Hochstein, 1986). Other potential electron acceptors used by many species are dimethylsulfoxide, trimethylamine N-oxide (Oren and Trüper, 1990) and fumarate (Oren, 1991). Halobacterium salinarum, but none of the many other aerobic halophilic Archaea tested (Oren and Litchfield, 1999), is able to grow fermentatively on L-arginine (Hartmann et al., 1980). However, the group of halophilic microorganisms that have specialized in anaerobic fermentative growth is that of the Haloanaerobiales (low G+C branch of the Gram-positive Bacteria) (Oren, 1992). Different sugars and in some cases also amino acids are fermented to products such as acetate, ethanol, butyrate, hydrogen, and carbon dioxide (Lowe et al., 1993; Mermelstein and Zeikus, 1998; Oren, 1992; Rainey et al., 1995). In low-salt anaerobic environments breakdown of organic compounds is completed by the cooperative action of a variety of microbial processes, including fermentation, dissimilatory sulfate reduction, methanogenesis, and possibly also activity of proton-reducing acetogens that degrade compounds such as ethanol, butyrate, and others to hydrogen and acetate. Not all these processes have been identified as yet in anaerobic hypersaline environments (Oren, 1988b). For example, no reports on the occurrence and activity of proton-reducing acetogens in hypersaline environments have been published as yet. This lack of information may be due to the difficulty in handling these intriguing bacteria, which are being studied by a very small number of microbiologists only. Thus any claim that such halophilic organisms do not occur in nature may be premature. Dissimilatory sulfate reduction occurs up to quite high salt concentrations. Black, sulfidecontaining sediments are often found on the bottom of salt lakes and saltern ponds almost up to NaCl saturation. A number of halophilic sulfate reducers have been isolated in recent years. The most salt-tolerant isolate thus far is Desulfohalobium retbaense, isolated from Lake Retba in Senegal, which was documented to grow at NaCl concentrations of up to 24% (Ollivier et al., 1991). Other halophilic isolates such as Desulfovibrio halophilus and Desulfovibrio oxyclinae tolerate NaCl concentrations of up to 18– 22.5% only (Caumette, 1993; Caumette et al., 1991; Krekeler et al., 1997; Ollivier et al., 1994). Most halophilic and halotolerant sulfate reducers isolated are incomplete oxidizers that grow

Life at High Salt Concentrations

269

on lactate and produce acetate. Only very recently was the first halophilic acetate-oxidizing sulfate-reducing bacterium isolated: Desulfobacter halotolerans was obtained from the bottom sediments of the Great Salt Lake (Brandt and Ingvorsen, 1997). This organism has a rather restricted salt range, being unable to grow above 13% NaCl. Possibly, bioenergetic constraints define the upper salinity limit at which the different dissimilatory processes can occur (Oren, unpublished data). The sulfate reducers are Proteobacteria that use organic compatible solutes to provide osmotic balance, a strategy that is energetically much more expensive than the use of inorganic ions for that purpose (see below). Accumulation of trehalose and glycine betaine was documented in Desulfovibriohalophilus (Welsh et al., 1996). Dissimilatory sulfate reduction provides relatively little energy, and therefore the need to spend a substantial part of the available energy for the production of organic osmotic solutes may set the upper limit to the salt concentration at which these bacteria can grow. The oxidation of lactate to acetate and CO2 yields much more energy (2 Lactate- + SO42- Æ 2 Acetate- + 2HCO3- + HS- + H+ ; DGo¢ = -160.1 kJ) than the oxidation of acetate with sulfate as electron acceptor (Acetate- + SO42- Æ 2HCO3+ HS-; DGo¢ = -47.7 kJ). This difference may possibly explain the apparent lack of complete oxidizers at the highest salt concentration range. The main methanogenic processes in freshwater environments are the reduction of CO2 with hydrogen and the aceticlastic split. Neither of these reactions has been shown to occur at high salt concentrations. Solar Lake (Sinai) sediments (70–74 g/liter salt) did not show any methanogenesis from acetate or from H2 + CO2 (Giani et al., 1984). The highest salt concentration at which methanogenesis from H2 + CO2 was demonstrated in nature was 88 g/liter (Mono Lake, CA) (Oremland and King, 1989). The most halotolerant isolate that grows on H2 + CO2 is Methanocalculus halotolerans obtained from an oil well. This organism grows up to 12% NaCl with an optimum at 5% (Ollivier et al., 1998). The upper salinity boundary for the use of acetate as methanogenic substrate is probably even lower, but available data are few. To my knowledge, no cultures of aceticlastic methanogens are extant that grow above 4–5% NaCl. Energetic constraints may explain the apparent lack of truly halophilic methanogens that grow on H2 + CO2 or on acetate. In contrast to the aerobic halophilic Archaea of the order Halobacteriales which contain inorganic ions for osmotic stabilization, the methanogens use the energetically more expensive option of synthesizing organic osmotic solutes (see Table 5). The aceticlastic split yields very little energy

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(DGo¢ = -31.1 kJ per mol acetate). The free energy yield during growth on hydrogen is -34 kJ per mol of hydrogen, not much higher than that on acetate. Methanogenesis does occur, however, at much higher salt concentrations. The most salt-tolerant methanogens known in culture are Methanohalobium evestigatum and Methanohalophilus portucalensis, which grow in up to 25–26% NaCl (Boone et al., 1993; Lai and Gunsalus, 1992; Zhilina and Zavarzin, 1987). Additional moderately halophilic methanogens have been isolated, growing optimally at 4–12% salt (e.g., Methanohalophilus mahii, Methanohalophilus halophilus, Methanohalophilus portocalensis, and Methanohalophilus zhilinae). The energy sources used by these methanogens are methylated amines, methanol, and dimethylsulfide (Oremland and King, 1989; Zhilina and Zavarzin, 1987, 1990; see also the review paper by Ollivier et al., 1994). The substrates, such as trimethylamine and dimethylsulfide, used by these bacteria in their natural environment are largely derived from microbial degradation of methylated compounds that serve as organic osmotic solutes in many halophilic microorganisms (Oremland and King, 1989; Zhilina and Zavarzin, 1990; see also Table 5). Thermodynamic calculations show that the energy yield on methylated amines is relatively large (between -92 and -191 kJ per mol of substrate transformed), and this may explain, at least in part, why growth of methanogenic Archaea on methylated amines may occur up to high salt concentrations. While, as discussed above, methanogens growing on H2 + CO2 appear to be absent in hypersaline environments, halophilic homoacetogenic bacteria that use the same substrates for the production of acetate have been isolated (Zavarzin et al., 1994; Zhilina and Zavarzin, 1990; Zhilina et al., 1996). Acetohalobium arabaticum is able to grow between 10–25% NaCl with an optimum at 15–18% (Zhilina and Zavarzin, 1990). At first sight, reaction thermodynamics do not explain why halophilic homoacetogenic bacteria do occur when CO2-reducing methanogens do not, as the acetogenic reaction yields even less energy than the methanogenic reaction (-26.1 kJ and -31.1 kJ per hydrogen oxidized, respectively). However, the halophilic homoacetogens belong to the order Haloanaerobiales (Rainey et al., 1995; Zhilina et al., 1996), a group that uses the energetically cheaper option of accumulating inorganic ions to establish osmotic balance (Oren, 1986; Oren et al., 1997; Rengpipat et al., 1988). Halophilic aerobic chemoautotrophic bacteria that obtain their energy from the oxidation of reduced sulfur compounds are known. Thiobacillus halophilus, isolated from a hypersaline lake in Western Australia, grows in as much as

CHAPTER 1.9

24% NaCl (Wood and Kelly, 1991). However, autotrophic oxidation of NH4+ to NO2- was never demonstrated above 150 g/liter salt, and the salt limit for the oxidation of NO2- to NO3- may be even lower (Rubentschik, 1929). To my knowledge no halophilic or halotolerant ammonia- or nitrite-oxidizing bacteria are extant in culture that are able to grow at salinities significantly exceeding those of seawater. Nitrosococcus halophilus, with an optimum at 4% NaCl and a maximum at 9.4% may be the most halophilic strain isolated to date (Koops et al., 1996). An attempt to demonstrate nitrification in a microcosm simulation of the microbiology of the Great Salt Lake at a total salt concentration above 30% yielded negative results (Post and Stube, 1988). Lack of energy source is probably not the main reason: ammonia, and not nitrate, is the dominant inorganic nitrogen species in most or all hypersaline water bodies, and it generally occurs in quite high concentrations, in the Dead Sea even in the millimolar range. Energetic constraints may be the cause for the apparent lack of halophilic nitrifying bacteria in nature, as only very small amounts of energy are gained from the oxidation of ammonia and of nitrite. Thermodynamic constraints cannot explain the apparent lack of aerobic methane oxidation in hypersaline environments. Methane oxidation is a highly exergonic process (CH4 + 2O2 Æ HCO3+ H+ + H2O; DGo¢ = -813.1 kJ). However, even in an environment with a relatively low salinity such as the epilimnion of Solar Lake, Sinai, during winter stratification (about 9% salt), no methane oxidation could be measured in spite of the availability of both methane and oxygen (Conrad et al., 1995). Recent reports on the occurrence of methane oxidation in sediments of hypersaline reservoirs in Ukraina and Tuva (up to 330 g/liter total dissolved salts) and on the isolation of halophilic methanotrophs from these environments (Kalyuzhnaya et al., 1998; Khmelenina et al., 1996, 1997; Sokolov and Trotsenko 1995) indicate that the existence of halophilic methane oxidizers is at least thermodynamically feasible, and that the earlier reported lack of methane oxidation in other hypersaline environments (Conrad et al., 1995; Slobodkin and Zavarzin, 1992) should have other reasons.

Physiological and Biochemical Properties of Halophilic Prokaryotes As biological membranes are permeable to water, any microorganism living at high salt concentrations has to maintain its intracellular environment at least isoosmotic with the salt concentration in its environment, and even

CHAPTER 1.9

Life at High Salt Concentrations

hyperosmotic when a turgor pressure has to be maintained (Brown, 1976, 1990; Csonka, 1989; Vreeland, 1987). Two fundamentally different strategies exist that enable halophilic and halotolerant prokaryotes to cope with the osmotic stress exerted by the high ionic strength of their hypersaline environment. The first option, used by the aerobic Archaea of the order Halobacteriales and by the anaerobic Bacteria of the order Haloanaerobiales, is based on the accumulation of high concentrations of inorganic ions in the cytoplasm. In most cases, K+ rather than Na+ is the dominant intracellular cation, and Cl- is the dominant anion. Presence of molar concentrations of inorganic ions requires special adaptations of the entire intracellular enzymatic machinery. The “salt-in” strategy permits little flexibility and adaptability to changing conditions, as many salt-adapted enzymes and structural proteins require the continuous presence of high salt for activity and stability. The adaptive evolution of proteins and salinity-mediated selection of their properties has recently been reviewed (Dennis and Shimmin, 1997). The second strategy is to prevent high salt concentrations from reaching the cytoplasm, and maintaining “conventional” enzymes and other proteins, not specifically designed to function at high ionic strength. Low intracellular ionic concentrations are maintained by active pumping of ions out of the cells. Osmotic equilibrium is provided by organic solutes that are either produced by the cells or accumulated from the medium (Kempf and Bremer, 1998). Such “compatible” solutes are low-molecular-weight organic compounds, soluble in water at high concentrations, and not inhibitory to enzymatic activities even in the molar concentration range. The intracellular concentrations of the organic solutes are regulated according to the salinity of the external medium. Thus, the use of organic osmotic solutes provides a great deal of flexibility and adaptability to an often, wide range of

271

salt concentrations, with the possibility of rapid adaptation to changes in the salinity of the medium. The strategy of maintaining isoosmotic concentrations of organic osmotic solutes is used by most halophilic and halotolerant Bacteria (with the exception of the Haloanaerobiales, as stated above) and by the halophilic methanogenic Archaea. Halophilic eukaryotic microorganisms also use organic compatible solutes for osmotic stabilization. Under certain conditions, the alkaliphilic members of the Halobacteriales also make use of an organic osmoticum (2-sulfotrehalose) to aid in the achievement of osmotic equilibrium with the environment (Desmarais et al., 1997).

The “Salt-In” Strategy Analyses of intracellular ionic concentrations in different aerobic halophilic Archaea show that these microorganisms maintain extremely high salt concentrations inside their cells. Moreover, the ionic composition of their intracellular milieu differs greatly from that of the outside medium, with K+ being the main intracellular cation (Table 2). The representatives of the order Haloanaerobiales (low G+C Gram-positive branch of the Bacteria) display a number of physiological and biochemical properties that are characteristic for the halophilic aerobic Archaea, rather than for the moderately halophilic aerobic Bacteria which use the organic solute strategy. No organic osmotic solutes have been found as yet in this group of anaerobic halophilic fermentative Bacteria (Mermelstein and Zeikus, 1998; Oren, 1986; Oren et al., 1997; Rengpipat et al., 1988). High concentrations of Na+, K+ and Cl- were measured inside the cells of Haloanaerobium praevalens, Haloanaerobium acetoethylicum, and Halobacteroides halobius, high enough to be at least isotonic with the medium (Table 3). In exponen-

Table 2. Estimates of intracellular ionic concentrations in aerobic halophilic Archaea of the order Halobacteriales. Medium concentration +

Species

Na

Halobacterium salinarum Halobacterium salinaruma Halobacterium salinaruma Haloarcula marismortuib Haloarcula marismortuia Haloarcula marismortuic Halococcus morrhuae

4.0 3.7 3.33 3.9 3.9 3.9 4.0

a

K

+

0.032 0.013 0.05 0.004–0.007 0.001–0.004 0.0075– 0.032

Mg

2+

0.1 0.13 0.15 0.15 0.15

Intracellular concentration Cl

−

3.9 3.9 3.9

Na

+

1.37 1.63 0.80 1.2–3.0 1.6–2.1 0.5–0.7 3.17

K+ 4.57 2.94 5.32 3.77–5.5 3.7–4.0 3.7–4.0 2.03

Mg2+

Cl− 3.61

0.12

2.3–4.2 3.2–4.1 2.3–2.9 3.66

Late exponential growth phase cells; bEarly exponential growth phase cells; cStationary growth phase cells. For additional information see text. Data were derived from Christian and Waltho (1962), Ginzburg et al. (1970), Lanyi and Silverman (1972), and Matheson et al. (1976). All concentrations are in molar units, except those relating to Haloarcula marismortui, which are expressed in molal units.

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CHAPTER 1.9

Table 3. Intracellular ionic concentrations of halophilic anaerobic Bacteria of the order Haloanaerobiales. Medium concentration

Intracellular concentration

Species

Na+

K+

Cl−

Na+

K+

Cl−

Haloanaerobium praevalens

0.99 2.22 2.22 3.08 1.16 2.52 1.56

0.013 0.013 0.013 0.013 0.032 0.034 0.013

1.07 2.30 2.30 3.16 1.40 2.70

0.44 1.52 0.44 2.63 0.92 1.50 0.54

0.96 1.59 1.14 2.05 0.24 0.78 0.92

2.24 1.26 3.28 1.20 2.50

Haloanaerobium praevalensa Haloanaerobium acetoethylicum Halobacteroides halobius a

Data obtained by X-ray microanalysis in the electron microscope. Values probably underestimate the true values. Data were derived from Oren (1986), Oren et al. (1997), and Rengpipat et al. (1988).

tially growing cells, K+ was the major cation. Stationary phase cells showed a high variability among individual cells, part of the cells containing high concentrations of NaCl rather than of KCl (Oren et al., 1997). The huge potassium concentration gradient over the cytoplasmic membrane (often up to three orders of magnitude) and also the generally large sodium gradient present can be created and maintained only at the expense of energy. Also the chloride ion is far from thermodynamic equilibrium, as the presence of an inside negative membrane potential would tend to expel Clfrom the cell. The peculiar ionic composition of the cells’ cytoplasm and the concentration gradients over the cell membrane are the result of the cooperative action of different ion pumps, antiporters, and other transport proteins. The most important ones are summarized in Fig. 9, and the numbers in square brackets in the explanation below refer to the different parts of that figure. hv

+ –

2

H+

ATP H+ +Pi

ATP

–

Cl–

3

1

+

hv

H+

8 Na+ 7 Na+ Cl–

– +

6 K+

5

4

H+

Amino Na+ acids

Fig. 9. Ion movements in the aerobic halophilic Archaea (order Halobacteriales): [1] proton extrusion via respiratory electron transport; [2] light-driven proton extrusion mediated by bacteriorhodopsin; [3] ATP formation by ATP synthase, driven by the proton gradient. Alternatively, this system can serve to generate a proton gradient at the expense of ATP during fermentative growth on arginine. [4] electrogenic sodium/proton antiporter; [5] sodium gradient-driven inward amino acids transport; [6] potassium uniport, driven by the membrane potential; [7] light-independent chloride transport system, probably coupled with inward transport of sodium. [8] halorhodopsin, the primary, light-driven chloride pump. For details see text.

In the Halobacteriales, respiratory electron transport with oxygen or other electron acceptors is accompanied by the extrusion of protons [1], generating a primary proton electrochemical gradient (acidic outside, alkaline inside, positive outside, negative inside). Those species that contain the retinal protein bacteriorhodopsin in their membranes may also use light energy for the direct generation of the proton electrochemical gradient [2]. The primary proton gradient is the driving force for all energy-requiring processes within the cell. Thus, ATP formation is mediated by the membrane-bound ATP synthase that couples phosphorylation of ADP with an inward flux of H+ [3]. The membrane ATP synthase may also be used in the reverse direction, the build-up of a proton electrochemical gradient at the expense of ATP. This process is relevant in cases in which ATP formation by substrate-level phosphorylation is the primary energy-yielding process in the cell. This is the case (e.g., in Halobacterium salinarum) when growing anaerobically by fermentation of arginine (Hartmann et al., 1980), or in the anaerobic Bacteria of the order Haloanaerobiales, which obtain their energy by fermentation of sugars or amino acids (Oren, 1992). The membranes of all halophiles investigated possess high activities of Na+/H+ antiporters, which use the proton electrochemical gradient as the driving force for the extrusion of Na+ from the cell [4] (Hamaide et al., 1983; Lanyi and MacDonald, 1976; Luisi et al., 1980). In Halobacterium salinarum, the antiporter was shown to be electrogenic and probably has a stoichiometry of 2 H+/Na+ (Lanyi and Silverman, 1979). In addition to its function of keeping intracellular Na+ concentrations at the desired low levels, the Na+/ H+ antiporter activity can be expected to play an important role in the regulation of the intracellular pH. The sodium gradient thus established can in its turn be used to drive certain endergonic processes. Thus, many of the membrane transport systems for amino acids and other compounds in the aerobic halophilic Archaea are energized by

CHAPTER 1.9

cotransport with Na+ ions [5]. The same is true for many moderately halophilic Bacteria, which also maintain a relatively low intracellular Na+ concentration (Shindler et al., 1977; Ventosa et al., 1998). The Na+ gradient thus serves to some extent as an energy reserve. It is generally accepted that the negative inside membrane potential is the driving force for the massive K+ accumulation. The membranes of halophilic Archaea were found to be highly permeable to potassium. K+ ions probably enter the cells via a uniport system in response to the membrane potential (Wagner et al., 1978) [6]. K+ enters the cell as Na+ is ejected by the electrogenic Na+/H+ antiporter, thus maintaining electroneutrality. Evidence for such a mechanism was found in experiments showing accumulation of radioactively labeled rubidium (a potassium analog) ions in right-side-out vesicles of Halobacterium salinarum as a reaction to sodium extrusion, following excitation of the light-driven primary proton pump bacteriorhodopsin (Kanner and Racker, 1975). In addition, a K+ transport system analogous to the Kdp system (the P-type K+-translocating ATPase) of Escherichia coli was detected in Haloferax volcanii (Meury and Kohiyama, 1989). This system requires ATP for activation. The high internal Cl- concentration is not in equilibrium with the large negative-inside electrical potential that accompanies the H+ circulation and the Na+ efflux. Thus, electrical potential-driven passive chloride movement can result only in a loss of chloride from the cells rather than in the required uptake. An increase in the amount of intracellular Cl- is essential if the cells should increase their volume during growth and cell division. It has been suggested that during growth the net flux of ions should result in K+ uptake and excess Na+ loss, and that Cl- uptake should be equal to the difference, and thereby provide a net gain of intracellular KCl commensurate with the gain in intracellular volume (Lanyi, 1986; Schobert and Lanyi, 1982). Two energy-dependent inward chloride pumps have been identified in Halobacterium cells. The first is a light-independent transport system, which is probably driven by symport with Na+ (Duschl and Wagner, 1986) [7]. The second is light-driven, and is based on the retinal protein halorhodopsin, a primary inward Cl- pump present in Halobacterium salinarum, in Natronomonas pharaonis, and probably also in additional halophilic Archaea (Lanyi, 1986; Schobert and Lanyi, 1982) [8]. The presence of molar concentrations of salts is generally devastating to proteins and other macromolecules. It causes aggregation or collapse of the protein structure by enhancing hydrophobic interactions within and between

Life at High Salt Concentrations

273

protein molecules and interferes with essential electrostatic interactions within or between macromolecules by charge shielding. And because of salt ion hydration, it reduces the availability of free water below the level required to sustain essential biological processes (Dennis and Shimmin, 1997; Zaccai and Eisenberg, 1991). The presence of high intracellular salt concentrations thus requires special adaptations of the whole enzymatic machinery of the cell. Cells thus adapted are able to function in the presence of high salt. However, these adaptations make the cells strictly dependent on the continuous presence of high salt concentrations for the maintenance of structural integrity and viability (Ebel et al., 1999; Eisenberg, 1995; Eisenberg and Wachtel, 1987; Eisenberg et al., 1992; Lanyi, 1974). As a result, the aerobic halophilic Archaea display little flexibility and adaptability to changes in the external salt concentration. Most enzymes and other proteins of the Halobacteriales denature when suspended in solutions containing less than 1–2 M salt. Many enzymes are more active in the presence of KCl than of NaCl, agreeing well with the finding that K+ is intracellularly the dominating cation. “Salting-out” salts stabilize, while “salting-in” salts inactivate halophilic enzymes. The behavior of different salts coincides with the lyotropic Hofmeister series (Lanyi, 1974). Similarly, intracellular enzymes from the fermentative anaerobic Bacteria (order Haloanaerobiales) generally function better in the presence of molar concentrations of salts than in salt-free medium, and they can be expected to be fully active at the actual salt concentrations present in the cytoplasm (Oren and Gurevich, 1993; Rengpipat et al., 1988; Zavarzin et al., 1994). Most proteins of the Halobacteriales contain a large excess of the acidic amino acids, glutamate and aspartate, and few basic amino acids, lysine and arginine. The high content of acidic side groups was first recognized during analyses of the bulk protein of Halobacterium and Halococcus (Reistad, 1970). The malate dehydrogenase of Haloarcula marismortui has a 10.4 mol% excess of acidic residues and the cell envelope glycoprotein of Halobacterium salinarum even 19–20 mol%. Owing to the high acidity of the proteins of the halophilic Archaea, isoelectric focusing is of little use for protein characterization and isolation. In Halococcus salifodinae, all proteins were found to have isoelectric points between 3.8 and 4.5 (Denner et al., 1994). The bulk cellular protein of the members of the Haloanaerobiales tested is also highly acidic (Oren, 1986). It has been argued that the excess of acidic residues may be a major factor determining the halophilic character of the proteins: excess of

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negative charges on the protein surface makes the structure unstable because of the mutual repulsion of the side groups. Only when high concentrations of cations are added to shield the negative charges can the protein maintain its proper conformation required for structural stability and enzymatic activity. Shielding of negative charges by cations undoubtedly plays an important part in the effects of salt on the enzymes and other proteins of the halophiles. A theoretical analysis of the contribution of electrostatic interactions in Haloarcula marismortui ferredoxin and malate dehydrogenase shows that the repulsive interactions between the acidic residues at the protein surface are a major factor in the destabilization of halophilic proteins in low-salt conditions (Elcock and McCammon, 1998). However, Lanyi (1974) and Lanyi and Stevenson (1970) stated that all the effects of salts cannot be due to charge-shielding action alone, as the concentrations required are too high. Maximal electrostatic-charge shielding would be achieved already in about 0.1 M salt or 0.5 M at most, and in even much lower concentrations of divalent cations. However, a high content especially of glutamate may be favorable, as glutamate has the greatest water binding ability of any amino acid residue. This may have important implications when considering the need of any functional protein to maintain a proper hydration shell. Another prominent feature of the proteins of the Halobacteriales is their low content of hydrophobic amino acid residues, generally offset by an increased content of the borderline hydrophobic amino acids, serine and threonine (Lanyi, 1974). The requirement for extremely high salt concentrations for structural stability of the proteins can probably to a large extent be attributed to the low content of hydrophobic residues and the accordingly weak hydrophobic interactions within the protein molecules. High salt is then needed to maintain the weak hydrophobic interactions. Entropy increases when non-polar groups turn away from the water phase and interact with each other to form hydrophobic interactions. These interactions seem to be driven more by an avoidance of water than by an active attraction between the non-polar molecules (Lanyi, 1974). At higher salt concentrations, new hydrophobic interactions are formed having insufficient stability in water, and the molecule assumes a more tightly folded conformation. The possible involvement of the weak hydrophobic interactions in the salt requirement of the halophilic proteins is supported by the finding that certain enzymes from halophilic Archaea (including threonine deaminase, aspartate carbamoyltransferase, and alanine dehydrogenase) show cold lability: their maximal

CHAPTER 1.9

stability is reached at temperatures greater than 0∞C and decreases at lower temperatures. The effect may be considered in terms of water structure: at lower temperature the size of the cluster of water molecules is increased, and hydrophobic groups can interact more easily, breaking the hydrophobic interactions (Lanyi, 1974). Detailed studies of the malate dehydrogenase of Haloarcula marismortui have contributed much valuable information on the possible mechanisms involved in the halophilic behavior of proteins. Techniques such as velocity sedimentation, light scattering, neutron scattering, and circular dichroism measurements have been used to obtain information on the structural changes occurring as a function of changing salt concentrations and the hydration properties of the protein (Eisenberg, 1995; Eisenberg and Wachtel, 1987; Mevarech and Neumann, 1977; Pundak and Eisenberg, 1981; Pundak et al., 1981). These studies have shown that the halophilic properties of the enzyme are related to its capacity of associating with unusually high amounts of salts, and led to the formulation of a thermodynamic “solvation-stabilization model,” in which the halophilic protein has adapted to bind hydrated ions cooperatively via a network of acidic groups on its surface (Ebel et al., 1999). X-ray diffraction studies on crystals of the halophilic malate dehydrogenase and the ferredoxin of Haloarcula marismortui and the dihydrofolate reductase of Haloferax volcanii have added much important information (Dym et al., 1995; Frolow et al., 1996; Pieper et al., 1998). These studies showed how the carboxylic groups on the acidic residues are used to sequester, organize, and arrange a tight network of water and hydrated K+ ions at the surface of the protein, and how an unusually large number of internal salt bridges between strategically located basic amino acid residues are formed to give the protein its internal structural rigidity. These salt bridges appear to be important determinants in the stabilization of the threedimensional structure of halophilic proteins. Intervening solvent molecules shield the negative charges of the carboxylic acid groups on the protein surface from each other. Comparison of the Haloarcula marismortui ferredoxin with the plant-type 2Fe-2S ferredoxin showed that the surface of the halophilic protein is coated with acidic residues, except for the vicinity of the iron-sulfur cluster, and that the halophilic protein contains two additional helices near the N-terminus. These helices form a separate hyperacidic domain, postulated to provide extra surface carboxylates for solvation. Bound water molecules on the protein surface have on the average 40% more hydrogen bonds than in a typical non-halophilic protein crystal

CHAPTER 1.9

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275

driven Na+ pumps detected in some moderate halophiles such as Salinivibrio costicola (Tokuda and Unemoto, 1983). Organic compatible solutes make up the major part of the osmotically active compounds in the cells’ cytoplasm. Compatible solutes are polar, highly soluble molecules, most of them uncharged or zwitterionic at the physiological pH. The list of compounds known to be synthesized as compatible solutes by halophilic microorganisms is steadily growing (Galinski, 1993, 1995; Galinski and Trüper, 1994; Reed, 1986; Trüper et al., 1991; Ventosa et al., 1998; Wohlfarth et al., 1990). Figure 10 shows the main osmotic solutes identified thus far in prokaryotes, and Table 5 provides information on the taxonomic groups in which the different solutes have been detected. The accumulation of “compatible” osmotic solutes achieves osmotic equilibrium while still enabling activity of “conventional”, non-saltadapted enzymes (Galinski, 1993, 1995). Many prokaryotic cells contain cocktails of different compatible solutes rather than relying on a single compound (Galinski, 1995). The concentrations of the osmotic solutes are regulated according to the salt concentration in which the cells are found (Galinski and Louis, 1999), and can be rapidly adjusted as required when the outside salinity is changed (by synthesis or uptake from the medium upon salt upshock; by degradation or transformation into osmotically inactive forms; or by excretion following dilution stress) (Trüper and Galinski, 1990). The use of organic osmotic solutes thus bestows a high degree of flexibility and adaptability. Compatible solutes are strong, water structure formers and as such they are probably excluded from the hydration shell of proteins. This “preferential exclusion” probably explains their function as effective stabilizers of the hydration shell of proteins. This phenomenon of nonspecific exclusion is often described in terms of increased

structure. These water molecules are thus tightly bound within the hydration shell by proteinwater and water-water hydrogen bonds and by hydration of interspersed K+ ions (Frolow et al., 1996). A recent study of the glutamate dehydrogenase of Halobacterium salinarum showed the surface of the molecule being covered with acidic residues and displaying a significant reduction in exposed hydrophobic character as compared to non-halophilic counterparts. The low lysine content helps to increase the overall negative charge on the protein surface but also serves to decrease the hydrophobic fraction of the solvent-accessible surface (Britton et al., 1998).

The “Low Salt-In” Strategy The second option, realized in most halophilic and halotolerant representatives of the Bacteria and also in the halophilic methanogenic Archaea, involves the maintenance of a cytoplasm much lower in salt than the outside medium. Table 4 summarizes estimates of intracellular salt concentrations in a number of aerobic halophilic Bacteria. While in some cases the apparent intracellular ionic concentrations are in the molar range (possibly in part due to technical difficulties related to the exact assessment of the intracellular water volumes in cell pellets) (Ventosa et al., 1998), it is clear that the intracellular salt concentrations are generally insufficient to provide osmotic balance. Generally the intracellular Na+ concentrations are kept low. Outwarddirected sodium transporters in the cytoplasmic membrane (in most cases electrogenic Na+/H+ antiporters) are highly important both in maintaining the proper intracellular ionic environment and in pH regulation (Hamaide et al., 1983; Ventosa et al., 1998). Outward-directed Na+ transporters may include primary respiration-

Table 4. Intracellular ionic concentrations of selected aerobic halophilic Bacteria. Medium concentration +

Species

Na

Halomonas elongata

1.38 3.4 4.4 1.0 3.0 2.0 3.0 2.0

Halomonas canadensis Halomonas halodenitrificans “Pseudomonas halosaccharolytica” Salinivibrio costicola

K

+

0.02 0.01 0.04 0.04 0.006 0.006 0.008

Intracellular concentration Cl

−

1.0 3.0 2.0 3.0

Na

+

0.31 0.63 0.62 0.31 1.07 1.15 1.04 0.90

K+ 0.02 0.02 0.58 0.47 0.12 0.89 0.67 0.57

Cl−

0.055 0.98 0.70

All data relate to exponentially growing cells. Data were derived from Christian and Waltho (1962), Masui and Wada (1973), Matheson et al. (1976), Shindler et al. (1977), and Vreeland et al. (1983). For more extensive data, see Table 5 in Ventosa et al. (1998).

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CH3

COO-

CH2

+

N

CH3

CHAPTER 1.9 H N+

CH3

CH3

Glycine betaine

CH3

NH

COO-

NH

CH3

CH2

N+

CH3

OH

H

C

OH

O

CH3

CH2OH

OH

C

C

O

C H

H

O

CH3

CH2OH OH

C H

OH

CH2

Acetylchloline

O C

H

CH2OH

Glucosylglycerol

CH2OH

CH2OH OH

CH3

OH

β-Hydroxyectoine

Ectoine

CH2OH

CH2OH OH

+ CH3

COO-

CH2

+

Choline

OH

NH +

N

CH3

CH3

Dimethylglycine

NH

O

CH3

COO-

CH2

Mannitol

OH OH

CH2OH OH

OH O

CH2OH

OH

CH2OH OH

O

OH

O

OH O SO3HO

OH

CH2OH

O

O O

Sucrose

CH2OH

OH

O

CH2OH OH

OH

OH OH

OH

Trehalose CH2

CH2

–

COO

CH2

C

H

NH2 NH3+ CH2 C C O CH2 COO– H

H+ H

CH2

Proline

CH2

CH3

NH

CH2 CH2

NH3+ C COO– CH2 H

CH3

CH2

NH

C O

CH2

Nδ-acetylomithine O CH2

C CH3

NH

CH2

CH2 CH2

COO–

CH

Nε-acetyl-β-lysine

NH3

+

O C CH2

NH

CH2

CH2

C H

H

NH2 C O

NH2

NH3+ C COO–

Nε-acetyllysine CH2

CH2

CH2

C

CH2

β-Glutamine

O C

O COO–

CH

C

Glutamine

H

NH3+

O NH2

2-Sulfotrehalose

C

C NH

O NH2

NH2

Nα-carbamoylglutamine amide

O

O C

CH3 CH3

CH2

C

NH

CH2

C

CH2

CH2

COO–

Dimethylsulfoniopropionate

C H

S+ CH3

O C

O

NH2

Nα-acetylglutaminylglutamine amide Fig. 10. Organic osmotic solutes documented to occur in halophilic and halotolerant Bacteria and Archaea.

surface tension of water, with the presence of solutes affecting the forces of cohesion between water molecules, minimization of entropy, and reinforcement of the hydrophobic effect. Com-

patible solutes display a general stabilizing effect by preventing the unfolding and denaturation of proteins caused by heating, freezing, and drying (Galinski, 1993, 1995).

CHAPTER 1.9

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277

Table 5. Distribution of selected organic osmotic solutes within the bacterial and the archaeal domains. Solute Glycine betaine Dimethylglycine Choline, Acetylcholine Ectoine, Hydroxyectoine Proline Glutamine β-Glutamine Nε-acetyllysine Nδ-acetylornithine Nε-acetyl-β-lysine Nα-carbamoyl-glutamine amide Nα-acetylglutaminyl-glutamine amide Sucrose Trehalose 2-Sulfotrehalose Mannitol Glucosylglycerol Dimethylsulfonio-propionate

Distribution Cyanobacteria, Anoxygenic phototrophic bacteria; Methanogenic bacteria; Actinopolyspora halophila; Is taken up by many heterotrophic bacteria and used as osmotic solute. Methanogenic bacteria Lactobacillus plantarum Heterotrophic Proteobacteria of the γ-subdivision; Halorhodospira spp., Rhodobacter sulfidophilus; Micrococcus spp., many bacilli; Marinococcus spp., Sporosarcina halophila; Brevibacterium Bacilli; Planococcus citreus; Salinicoccus sp. Corynebacteria Methanogenic bacteria Halobacillus halophilus, other bacilli Halobacillus halophilus, other bacilli Methanogenic bacteria Ectothiorhodospira marismortui Halochromatium; Thiohalocapsa; Rhodopseudomonas sp.; Azospirillum brasilense; Rhizobium meliloti; Pseudomonas aeruginosa Cyanobacteria Cyanobacteria, Halorhodospira spp. Alkaliphilic members of the Halobacteriales Pseudomonas putida Cyanobacteria; Rhodobacter sulfidophilus; Pseudomonas mendocina Marine cyanobacteria

For additional information see e.g., Desmarais et al. (1997); Galinski (1993, 1995); Hagemann et al. (1999); Imhoff (1993); Oren (1999); Trüper et al. (1991); Ventosa et al. (1998); and Wohlfarth et al. (1990).

Concluding Remarks A comparison of the two strategies of adaptation to high salt concentrations (“salt-in” versus use of organic osmotic solutes) shows that the salt-in strategy is energetically much less costly than the synthesis of organic, compatible solutes (Oren, unpublished data). However, it requires a fargoing adaptation of the whole intracellular machinery to the presence of high ionic concentrations. This energetically relatively cheap solution of balancing “salt-out” with “salt-in” is not widely used in nature. Evolutionary processes toward such adaptation, as described by Dennis and Shimmin (1997), have led to the establishment of two specialized groups, the aerobic extremely halophilic Archaea (Tindall, 1992) and the fermentative obligatory anaerobic Bacteria (Oren, 1992). The use of organic compatible solutes allows much more flexibility with respect to the range of salt concentrations tolerated, and does not require a high degree of adaptation of the intracellular enzymes. The enzymes do not greatly differ from those of non-halophilic prokaryotes, although they may have a somewhat increased content of acidic amino acids (Gandbhir et al., 1995). Many taxonomic groups, displaying a great metabolic diversity, use this strategy. Thus, many of the dissimilatory processes identified in freshwater environments can also take place at

high salinity. Certain metabolic types, however, such as methanogenesis from H2 + CO2 or from acetate, autotrophic nitrification, and others, seem to be absent above 10–15% salt. It is tempting to speculate that it is the too high energetic cost connected with adaptation to life at the highest salt concentrations that has prevented the evolution of halophiles performing these reactions. In any case, also in hypersaline environments, the prokaryotes display an amazing diversity that is well worth being studied indepth on the level of community structure and metabolism, phylogenetic diversity, and the molecular mechanisms of their adaptation to high salt. Acknowledgments. I thank Carol D. Litchfield (George Mason University, Fairfax, VA) for her helpful comments on the manuscript.

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CHAPTER 1.9 Zhilina, T. N., and G. A. Zavarzin. 1994. Alkaliphilic anaerobic community at pH 10. Curr. Microbiol. 29:109–112. Zhilina, T. N., G. A. Zavarzin, E. N. Detkova, and F. A. Rainey. 1996. Natroniella acetigena gen. nov. sp. nov., an extremely haloalkaliphilic, homoacetic bacterium: a new member of Haloanaerobiales. Curr. Microbiol. 32:320– 326.

Prokaryotes (2006) 2:283–308 DOI: 10.1007/0-387-30742-7_10

CHAPTER 1.10 c i l i hp i l ak lA

se toyrakorP

Alkaliphilic Prokaryotes TERRY ANN KRULWICH

Introduction and Definitions Alkaliphilic prokaryotes offer a wealth of opportunities for the isolation of natural products that can be advantageously applied to diverse industrial processes (Horikoshi and Akiba, 1982; Horikoshi, 1991; Horikoshi, 1996). Alkaliphiles also offer a wealth of opportunities to understand the mechanisms by which organisms thrive at pH 10–11.5, often present with other environmental challenges, such as high salt and high temperature (Krulwich, 1995; Krulwich et al., 1998). The diversity of alkaliphiles, which occur among aerobic and anaerobic archaea and prokaryotes as well as eukaryotic fungi (Grant et al., 1990; Jones et al., 1994; Horikoshi, 1991), likely broadens their industrial application. All alkaliphiles confront protein structure and function problems that involve catalysis at the external pH and regulation of cytoplasmic pH. However, among different alkaliphile groups, such as anaerobes, aerobes, ammonium-requiring sulfate-reducing thermophiles, halophiles, methylotrophs, methanogens, and photosynthetic bacteria, different approaches or features to the solution of these central as well as other groupspecific problems are likely. Alkaliphiles, an extraordinarily diverse group of bacteria that may have evolved over a very long period, provide us with the opportunity to probe their many adaptive strategies (Zavarzin, 1993; Duckworth et al., 1996; Jones et al., 1998). Also, understanding how alkaliphiles meet extreme challenges of pH homeostasis and other challenges can provide insights that are often translatable to the nonextremophile setting. Various definitions of an alkaliphile have been used. The only common standard is that the organism’s optimal pH for growth must be at least 2 units above neutrality (Kroll, 1990). To clearly distinguish between alkaliphiles and the more abundant alkaline-tolerant prokaryotes, most investigators of these organisms propose a higher pH standard and sometimes (Horikoshi, 1998) include an inability to grow well or grow at all at near-neutral pH values such as 6.5. As discussed below, growth at near-neutral pH may

be inversely related to how functional an alkaliphile is at the upper pH limit. The lower pH limit and pattern of growth over a broad pH range also depends upon the particular growth substrate (Gilmour and Krulwich, 1997; Krulwich et al., 1997) and on strain differences that are incompletely understood (Guffanti et al., 1986). My laboratory has used the term alkaliphile for bacteria that grow optimally on substrates (suitable for high pH growth) at pH values above 9.5. This value is chosen because it approximates the upper limit of cytoplasmic pH that is compatible with the growth of bacteria so far studied (Sturr et al., 1994; Krulwich et al., 1998). The term facultative alkaliphile has been applied to species and strains that are able to grow at both pH 6.5–7.5 and above 9.5, whereas alkaliphiles that grow only at 9.5 are termed obligate alkaliphiles.

Historical Notes Koki Horikoshi, a leading investigator of alkaliphiles, was the first to initiate broad-based studies of these bacteria, starting in the late 1960s. He has commented that when he began he found only sixteen prior literature references to alkaliphilic bacteria (Horikoshi, 1998). The earliest reports of bona fide alkaliphiles were those of Bacillus pasteurii by Gibson, 1934 and of Bacillus alcalophilus by Vedder, 1934. Investigators of physiology and protein structure have returned to these two alkaliphilic Bacillus species in recent years. In the early 1960s, Takahara and colleagues (Takahara et al., 1961; Takahara and Tanobe, 1960; Takahara and Tanobe, 1962) demonstrated that indigo dye reduction depends on maintaining sufficient alkalinity, and they improved the fermentation process by adding alkaliphilic Bacillus sp strain S-8, which they had isolated from an indigo ball undergoing fermentation at high pH. Over the years, Horikoshi and colleagues (Horikoshi, 1996) and others (e.g., van der Laan et al., 1991; Ito et al., 1998) advanced this tradition of optimizing alkaliphile fermentation processes or their products while

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also contributing to alkaliphile taxonomy and characterization. The diversity of alkaliphiles was subsequently extended by Grant and colleagues (Tindall et al., 1984; Grant et al., 1990; Grant et al., 1999; Jones et al., 1998), Zhilina and Zavarzin, 1994, and others who enumerated and identified bacteria and archaea that thrive in natural, selective environments such as the highly alkaline soda lakes in Africa and Asia. The number of known alkaliphiles and their diversity has vastly increased as a result of such systematic investigations. These studies also extended the earlier finding by (Kurono and Horikoshi, 1973; Horikoshi, 1991) that alkaliphiles require and thrive in added Na+, by showing that many categories of alkaliphiles are haloalkaliphiles. During the 1970s, my laboratory began to study energetic dilemmas of alkaliphiles, taking the chemiosmotic view of energy coupling found in bacterial and eukaryotic membrane systems (Mitchell, 1961). We studied the dilemma of how aerobic alkaliphilic bacteria can both extrude protons during respiration and achieve—as initially presumed and subsequently shown to be necessary—net acidification of the cytoplasm relative to the outside (Krulwich, 1995). As also noted by Garland, 1977, there was the further dilemma of what effect a “reversed pH gradient” would have on reducing chemiosmotic gradients (acid and cations out) and in turn on reducing the energetic driving force, which could create a problem with solute transport, flagellar rotation energization, and ATP synthesis. Perhaps some alkaliphiles generate compensatory electrical potentials across the membrane or use Na+ as a coupling ion, or perhaps some or all alkaliphiles had developed other interesting mechanisms for bioenergetic work under such circumstances. This area of study became and remains active in several laboratories (Krulwich and Guffanti, 1983; Krulwich and Guffanti, 1989; Krulwich and Ivey, 1990; Krulwich et al., 1998; Hirota and Imae, 1983; Sugiyama et al., 1985; Koyama et al., 1986; Koyama and Nosoh, 1995; Aono and Ohtani, 1990; Hamamoto et al., 1994). More recent investigations of alkaliphilic enzyme pH stability and optimum have drawn upon 3-dimensional crystal structures (Sobek et al., 1990; Sobek et al., 1992; van der Laan et al., 1992; Martin et al., 1997; Shirai et al., 1997) and properties that can be modeled or deduced from molecular characterizations (Teplyakov et al., 1992; van der Laan et al., 1996; Kobayashi et al., 1999). The physiology and study of bioenergetic properties of alkaliphiles also are enhanced by modern molecular biological techniques as well as proteome and genomic insights. Numerous alkaliphile genes have been sequenced, physical maps have been presented for three genes

CHAPTER 1.10

(Southerland et al., 1993; Park et al., 1994; Gronstad et al., 1998; Takami et al., 1999b), and the genome of an alkaliphilic BacillusM, Bacillus halodurans C-125, is currently being sequenced (Takami et al., 1999c). By consensus, there has been a shift in nomenclature from the use of the term “alkalophily” to use of the term “alkaliphily” during the 1990s. The more etymologically correct “alkali” has now become the convention.

Distribution and Taxonomic Groups Distribution Alkaliphiles are found in natural and industrial (or other man-made enrichments). In addition to the indigo dye process that has already been noted, sodium hydroxide has been used extensively in paper and pulp processing and calcium hydroxide in cement manufacture. Mining operations and certain food-processing activities also are settings for alkaliphile enrichment (Jones et al., 1998). Similarly, natural enrichments are diverse. Many of these, such as alkaline hot springs, are the source of interesting, generally alkaline-tolerant organisms but are insufficiently buffered to support the extraordinarily high pH values that are consistently maintained (Jones et al., 1998). On the other hand, naturally occurring soda lakes are stable, extremely alkaline environments (e.g., pH >11.5) that are widely distributed and typically found inland (e.g., soda lakes of the East African Rift Valley and of Central Asia). Their NaCl concentrations range from about 5% w/v to >15% w/v. The soda lakes have a paucity of calcium and magnesium ions because they are depressions formed from nonsedimentary rocks, and the sodium, chloride, and bicarbonate/carbonate are the dominant ions. The soda lakes often exhibit the pronounced color of organisms (e.g., cyanobacteria) that are the primary, photosynthetic actors in the nutrient cycle, and the hypersaline lakes often are the color of haloalkaliphiles. Both aerobic and anaerobic cycles occur (Jones et al., 1994; Jones et al., 1998). Alkaliphiles found in the soda lakes are diverse; some organisms are unique to these lakes and some (e.g. the fastidious haloalkaliphiles) appear to represent distinct lineages (McGenity and Grant, 1993; Jones et al., 1998; Grant et al., 1999). Also, evidence from fossil soda lakes, which are similar to those found today, suggests that these environments are of great antiquity. Together, these observations have led to suggestion that substantial evolution of many prokaryotesfound in this type of environment occurred in the soda lakes, i.e., these

CHAPTER 1.10

communities are very ancient sources of new species of bacteria (Zavarzin, 1993; Jones et al., 1998). If indeed many alkaliphiles evolved in natural soda lake enrichments, they must have spread beyond those boundaries. Some of the same alkaliphiles that are found in the ancient enrichments of the soda lakes are almost ubiquitous. They are present in garden soils (Guffanti et al., 1980, 1986) and in deep-sea trenches (Takami et al., 1999a) where the pH is not conducive to a thriving alkaliphile presence. Horikoshi and Akiba, 1982 note a substantial presence of alkaliphiles in soils of various pH values, albeit greater in alkaline soils.

Taxonomic Groups A summary of the taxonomic groups of prokaryotes isolated from soda lakes has been reproduced from a review by Jones et al., 1998 and presented at Table 1. The enormous taxonomic diversity of extreme alkaliphiles is evident and further reflected in the diversity of characteristics. Alkaliphilic cyanobacteria are among the primary photosynthetic organisms that produce oxygen; such organisms include Spirulina, Cyanospira, Synechococcus and Chorococcus. Anoxygenic phototrophic bacteria, such as Ectothiorhodospira, that use reduced sulfur compounds participate in the primary production via photosynthesis and also are part of the sulfur oxidizing limb of the sulfur cycle of the soda lakes. The sulfur cycle also includes aerobic sulfur oxidizing organisms (Sorokin et al., 1996) and anaerobic sulfate-reducing organisms such as Spirochaetes and Desulfonatronovibrio (Zhilina et al., 1996a; Zhilina et al., 1997). Bacillus species are among the most commonly found aerobic, eubacterial alkaliphiles both in soda lakes and in less selective environments (Horikoshi and Akiba, 1982; Krulwich and Guffanti, 1983; Guffanti et al., 1980; Guffanti et al., 1986; Takami et al., 1999a). Fritze et al., 1990, using DNA-DNA hybridization, and Nielsen et al. (Nielsen et al., 1994; Nielsen et al., 1995) using forty-seven physiological and biochemical characteristics as well as DNA base composition, hybridization, and 16S rDNA analyses, proposed clusters of alkaliphiles and alkaline-tolerant Bacillus. The 16S rDNA structure indicated two distinct groups within the Bacillus radiation (RNA groups 6 and 7) in which most alkaliphilic Bacillus isolates are found (Nielsen et al., 1994; Jones et al., 1998). Interestingly, the application of this comprehensive analytic approach resulted in grouping of strains that correlated roughly with somewhat distinct regions of the soda lake environment. B. alcalophilus and associated strains were mainly found

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in muds at the shoreline or dry regions of soda soil where organisms are subjected to fluctuating water levels and concomitant fluctuations in pH and salt levels; many of these strains require only low concentrations of Na+ for growth. The somewhat diverse “group 7” bacilli, related to Bacillus clarkii (Nielsen et al., 1994; Nielsen et al., 1995), are thought to be more prevalent in sediments and waters that are subject to less variability and these alkaliphiles typically exhibit requirements for higher Na+ for growth (Jones et al., 1998). The molecular physiological studies of Na+dependent pH homeostasis in alkaliphiles are consistent with such differences among strains (Krulwich et al., 1982; Garcia et al., 1983), and are beginning to identify respiratory chain components, transporters and cell surface molecules that may be of particular importance to extremophiles that face transitions or fluctuations (Hicks et al., 1991; Ito et al., 1997b). Some of the most intensely studied alkaliphilic Bacillus strains [i.e., Bacillus C-125 (Aono, 1995) and Bacillus firmus OF4 (Guffanti et al., 1986)] were characterized before the extensive matrix of physiological and molecular biological criteria, including 16S rDNA, were used to categorize them. In fact, their proposed species, Bacillus lentus and Bacillus firmus, respectively, were not included in the major alkaliphile clusters (Fritze, 1990; Nielsen et al., 1994; Nielsen et al., 1995). It was apparent that correlation of these alkaliphilic strains with their environmental patterns would require more precise placement. Therefore, recent work, including further biochemical tests and 16S rDNA sequencing, has resulted in proposals for reclassifying Bacillus C-125 from a probable Bacillus lentus strain to Bacillus halodurans C-125 (Takami and Horikoshi, 1999) and Bacillus firmus OF4 to Bacillus pseudofirmus OF4 (Takami and Krulwich, in press). The newly deduced relationships of these species to other Bacillus species are depicted in Figure 1. Clostridium strains as well as other diverse anaerobes are well represented in the prokaryotes of soda lakes (Table 1), but detailed studies or major applications of anaerobic, eubacterial alkaliphiles lag behind those of aerobic alkaliphiles. Perhaps the first described alkaliphilic anaerobe, a facultatively anaerobic strain that was subsequently classified as a new genus and species—Amphibacillus xylanus (Niimura et al., 1987; Niimura et al., 1989)—is the best studied. This interesting alkaliphile evokes possible shared themes with B. pasteurii, inasmuch as it depends upon high concentrations of ammonium for optimal growth. The cells possess an unusual glutamate dehydrogenase that may be involved in the assimilation of the ammonium (Jahns, 1996). A. xylanus is deficient in cytochromes, quinones and catalase (Niimura et al., 1987;

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CHAPTER 1.10 B.

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B. s NCDO1769 (X60646) B. DSM6307T (X76437) B. P (Z26923)

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Fig. 1. Unrooted phylogenetic tree showing the relationship of B. halodurans C-125 and B. pseudofirmus OF4 to other Bacillus strains. This figure is a slightly modified version of a figure presented by Takami and Krulwich, in press. The numbers indicate bootstrap samples, from among 1000, that supported the internal branches (Felsenstein, 1985). Bar = 0.01 Knuc unit.

Niimura et al., 1989), and possesses transporters that couple sugar or amino acid transport to Na+ gradients (i.e., they are Na+/solute symporters) and are markedly ammonium ion stimulated (Koyama, 1989; Koyama, 1993). Similarly, an apparently V-type NH4+- and Na+-stimulated ATPase has been described (Koyama, 1996; Kaieda et al., 1998). The mechanism and significance of the ammonium effect in this alkaliphile will be of considerable interest as will any special role ammonium might play in pH homeostasis. Because thermoalkaliphiles were not described in the earliest work on alkaliphiles, this combination of extreme adaptations was considered to be at first incompatible with life. However, during the past decade, major groups of at least moderately thermophilic alkaliphiles began to emerge. These include among others: a novel obligately alkaliphilic Clostridium species isolated from sewage (Li et al., 1993; Li et al., 1994; Wiegel, 1998); an asporogenous, Gram-positive ammonifying anaerobe from soda lake deposits, Tindallia magadii (Kevbrin et al., 1998); another xylan-degrading, anaerobic alkalithermophile, strain LB3A (Prowe et al., 1996; Sunna et al., 1997); and an actinomycete, Thermoactinomyces sp. strain HS682 (Tsuchiya et al., 1992). Stetter

Table 1. Taxonomic groups containing prokaryotes isolated from soda lakes (boldface type). Eubacteria Cyanobacteria Chroococcales Oscillatoriales Spirulina spp. Firmicutes (Gram-positive bacteria) Actinomycetes (high G+C Gram-positive bacteria) Actinomycetales Micrococcaceae Nocardiform actinomycetes Streptomyces Low G+C Gram-positive bacteria Bacillaceae Clostridiaceae Haloanaerobiales Proteobacteria Beta subdivision Delta subdivision Gamma subdivision Ectothiorhodospira Halomonadaceae Pseudomonas Spirochaetales Spirochaetaceae Spirochaeta Thermotogales Thermopallium Thermopallium natronophilum Archaea Euryarchaeota Halobacteriales Halobacteriaceae Halorubrum Halorubrum (Natronobacterium vacuolatum) Natrialba Natrialba (Natronobacterium) magadii Natronobacterium Natronobacterium gregoryi Unclassified Natronobacterium spp. Natronococcus Natronococcus amylolyticus Natronococcus accultus Unclassified Natronococcus spp. Natronomonas Natronomonas (Natronobacterium) phamonis Methanomicrobiales Methanosarcinaceae Methanohalophilus Methanohalophilus oregonensis Methanohalophilus zhilinaeae Methanohalophilus sp. Z-7936 Reproduced from Jones et al., 1998 with permission from the publisher.

and colleagues have even described a hyperthermophilic, alkaliphilic archaeum, Thermococcus alcaliphilus (Keller et al., 1995), completely laying to rest the notion of incompatibility of thermophily and alkaliphily. Conversely, Kimura and Horikoshi (Kimura and Horikoshi, 1989; Kimura and Horikoshi, 1990) have studied an

CHAPTER 1.10

alkalopsychrotrophic Micrococcus that produced an amylase that might be used in food processing. Finally, the haloalkaliphilic Archaea isolated from hypersaline lakes represent a burgeoning and fascinating group of multiple extremophiles that are now the subjects of broad-based studies. Originally such organisms were classified as Natronobacterium or Natronococcus (Tindall et al., 1984), halophiles that were most often responsible for the red color of the hypersaline soda lakes (Jones et al., 1998). More recently, the diversity of novel species and lineages has continued to grow (see Kanai et al., 1995; Zhilina et al., 1996b), including many from polymerase chain reaction (PCR)-based analyses of organisms that cannot be cultivated as yet (Grant et al., 1999). Proposals for reorganizing the taxonomy of this group have been made (Kamekura et al., 1997) in what is clearly a “moving target” situation. It is likely that the categories of alkaliphilic methylotrophs and methanogens, which already have been isolated in significant numbers and examples (Boone et al., 1993; Zhilina and Zavarzin, 1994), also will continue to increase in complexity (Kevbrin et al., 1997; Khmelenina et al., 1997).

Adaptation of the Proteins on the Outer Surface and Exoenzymes An initial set of generalizations describing the basis for protein stability under particular extreme conditions has begun to emerge from the results of experimental and modeling studies carried out on prokaryotic extremophile proteins from thermophiles (Chi et al., 1999; Elcock, 1998; Haney et al., 1999) and halophiles (Eisenberg et al., 1992; Elcock and McCammon, 1998). Generalizations that offer insight into the adaptation of alkaliphile proteins exposed to an extremely alkaline milieu are of similar interest. Many relevant observations have been reported, but broadly, applicable generalizations are not yet clearly in hand. Perhaps the reason lies with the proteins studied, which are often either thermophilic and/or halophilic or both, and therefore special subsets of highly alkaline enzymes or proteins. Or alkaliphily may mandate different responses by different functional categories of proteins. The deduced amino acid sequences, initially made available from genes that were cloned selectively from alkaliphile DNA libraries, revealed a significant number of examples in which the isoelectric point (pI) of the alkaliphile protein or protein domain was much lower than the homologues from other prokaryotes. These sequences had substantially increased aspartate and glutamate and strikingly reduced lysine and

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arginine. Examples included the prosequence (but not the mature form) of several alkaliphile proteases of the subtilisin type (van der Laan et al., 1991), SecY protein of B. halodurans C-125 (Kang et al., 1992), the “periplasmic” cytochrome c-binding domain of Cta (cytochrome oxidase) subunit II of B. pseudofirmus OF4 (Quirk et al., 1993), and a putative “periplasmic” loop of the FtsH protein from B. pseudofirmus OF4 (Ito et al., 1997a). The differences between this latter protein, as deduced from hydropathy analysis and sequence, and the homologue from Bacillus subtilis are shown in Fig. 2. Although not shown, there was no pattern of sequence difference in the two putative membrane spanning regions. The extensive pattern of added and substituted acidic residues and loss of basic ones is present throughout the loop, including regions very close to the membrane. In the Cta subunit, the first apparent substitutions of acidic residues for conserved basic ones are within 20 residues of the start of the large hydrophilic domain (Quirk et al., 1993); the net changes in the domain relative to non-alkaliphile homologues is shown in Table 2. The proximity of some of the substitutions to the membrane, in membrane-associated proteins, such as CtaC and FtsH, suggests that the selective pressure that leads to this type of change is exerted very close to the outer surface of the cytoplasmic membrane (Krulwich, 1995). These differences between alkaliphile and non-alkaliphile proteins or protein domains led to the suggestion that the alkaliphile might globally seek to minimize, in exposed proteins, the content of those basic residues that change charge over the range of external pH values to which the organism was regularly exposed (Krulwich et al., 1998). However, more complete and detailed information from both sequence and structural analyses of extracellular alkaliphile proteins and protein segments suggests that this explanation is unlikely and that the picture is more complex. A number of laboratories have begun to make chimeras of alkaliphile and non-alkaliphile homologues (Nakamura et al., 1991) to identify domains or residues involved in alkali-tolerance by selective mutagenesis and to design enzymes with increased alkaline-tolerance (Park et al., 1993). These studies give some indication that particular types of enzymes (for example, a particular domain such as a C-terminal domain in cellulases) can be associated with the alkaliphilic property (Nakamura et al., 1991). Moreover, there are now clear-cut examples of “periplasmic” loops and exoenzymes of alkaliphiles in which no evident substitution of acidic for basic residues occurs, relative to non-alkaliphile homologues. Indeed, some exoenzyme proteins,

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B. firmus OF4 VY Q R E P

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KD Q Y F L T H V P E G K G A D Q I F N A L K K DT V KV

Fig. 2. Comparison of the deduced sequence of the extracytoplasmic domain of the ftsH gene products of B. firmus OF4 (now B. pseudofirmus OF4) and B. subtilis. Acidic residues are in squares and basic residues are in circles. Reproduced with permission from the publisher from Ito et al. (1997a).

C-terminal

at least, show the opposite trend as will be discussed in detail in connection with subtilisin type proteases. Then how might the features of FtsH, CtaC, SecY and the prosequence of alkaline proteases be understood as an adaptation to high pH? Ikemura et al., 1987 and Chang et al., 1996 have suggested, based on findings from B. subtilis BPN and alkaliphilic Bacillus YaB protease studies, respectively, that the prosequence interacts (in trans, outside the cell) with the secreted but inactive protease to facilitate its folding to the active form. The highly charged nature of the prosequence (Ikemura et al., 1987), a feature that is conserved in alkaliphile and nonalkaliphile forms of the prosequence albeit with different sets of residues and vastly different isoelectric points, is proposed to effect this folding. van der Laan et al., 1991 accordingly argue that if acidic residues are not substituted for basic ones, then at the high pH values at which alkaline proteases function, the basic residues largely would be uncharged and protease activity compromised. By analogy the functions of other alkaliphile proteins that differ in the same way from non-alkaliphile homologues may involve

interactions that require the protein or domain partner to be highly charged. This suggestion is plausible for at least some of the examples already noted. The hydrophilic domain of CtaC must move in the “periplasm,” accepting an electron from complex III and then delivering it to CtaD. The chaperones and secretory proteins also must interact with various partners in specific ways that facilitate protection of conformation or movement. Following the expansion of this group of proteins will permit the adaptation to be more mechanistically elucidated. What about proteins without a special functional need for high charge density? The most intensive structural and modeling work to date on alkaliphile enzymes has focused on proteases, especially those of the subtilisin, serine protease type. This work has included X-ray crystallographic analysis, modeling and engineering of the active site (Sobek et al., 1990; Sobek et al., 1992; Teplyakov et al., 1992; van der Laan et al., 1992; van der Laan et al., 1996; Martin et al., 1997; Shirai et al., 1997). van der Laan et al., 1992 concluded that the 3-dimensional structure of the alkaline subtilisin-like protease PB92

Table 2. Comparisons of basic and acid amino acid contents of the hydrophilic cytochrome-c-binding domains of CtaC from alkaliphilic Bacillus pseudofirmus OF4 and non-alkaliphilic Bacillus cta-encoded cytochrome oxidases.1 Organism B. pseudofirmus OF4 B. subtilis B. stearothermophilus Bacillus PS3

Amino acids (#) in domain

Isoelectric point

Arginime + lysine

Aspartate + glutamate

GenBank Accession #

234 246 247 247

4.1 7.7 5.5 8.6

14 38 31 36

42 38 36 32

Q04441 P24011 BAA11111 Q03438

1 The hydrophilic domain starts at amino acid residue 109 from the N-terminal methionine or at amino acid residue 87 of the mature protein of the CtaC sequences; the homologous regions were identified from alignments with the domains from other strains.

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(also called SBA) revealed no “unique features” that could explain its very high pH optimum and suggested that sequence features could explain this optimum. In contrast to the observations on CtaC, FtsH, alkaline protease prosequences, and others already cited, PB92 and another highly alkaline protease, elastase YaB (Kaneko et al., 1989), had extremely high isoelectric points, as a result of decrease in aspartic acid residues and substitution of arginine for lysine residues, although not in equivalent positions. In addition, the number of tyrosines, which would be negatively charged at the enzyme’s pH optimum, is reduced (van der Laan et al., 1992). Interestingly, tyrosines do not always follow the pattern of glutamic and aspartic acid residues, i.e., increasing substantially in alkaliphilic proteins such as FtsH or consistently decreasing in high alkaline proteases (Shirai et al., 1997). It will be of interest, as more correlations become possible, to see whether the number of tyrosine residues, which may be “counted” as a likely source of negative charge predominantly in strongly obligate alkaliphiles, varies with particularly high pH optima for growth. Other alkaliphiles, whose pH range includes lower alkaline values at which tyrosine would be uncharged, might not adapt with a dependence upon this residue. In a report of the crystal structure of the alkaliphilic Bacillus M-protease, another subtilisintype serine protease, Shirai et al., 1997 further discussed the importance of sequence features in adaptation to alkaline conditions. They noted that of the three alkaliphile proteases of the subtilisin type with very high pH optima (PB92, M-protease, and elastase YaB) all had elevated isoelectric points relative to homologues with lower pH optima. M-protease had a markedly lower aspartic and glutamic acid content, which contributed to raising the isoelectric point. As noted in the other highly alkaline proteases, arginine was increased relative to lysine. The neutral hydrophilic amino acid residues, asparagine, glutamine and histidine, were increased, maintaining the solubility of the protein in water, perhaps by compensating for lost acidic residues and lysine. The increased arginine could contribute to the elevated isoelectric point and since arginine can retain a positive charge under more alkaline conditions than lysine, it is more suitable for ion pair formation. Ion pairs, including a significant number with arginine partners, increased. Hydrogen bonds also increased in the alkaliphile proteases relative to non-alkaliphile homologues. The investigators suggest that these features are important components of alkaline adaptation. Moreover, the “substituted residues” in the 3-dimensional structure had a biased distribution that correlated with peptide shifts

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Elastase YaB M-protease 0.60 PH92 0.81 4.52 A

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14.05 Alkaline B 12.81 93.77 9.28 17.77 Bacillopeptidase F Subtilisin Carlsberg

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Fig. 3. The phyogenetic tree of M-protease and related proteases showing evolutionary distances. The evolutionary distances are shown on the branches in percent between accepted point mutations. Two boxes indicate positions of hypothetical ancestors A and B (see Fig. 4). All of the branches are more than 95% probable from 1000 bootstrap reconstructions. This figure was reproduced from Shirai et al. (1997), with permission from the publisher.

hypothesized to be responsible for stabilizing conformation under alkaline conditions. Shirai et al., 1997 visualized the high alkaline subtilisin type proteases and the non-alkaliphilic ones as having branched off, respectively, from hypothetical ancestor proteases A and B (see Figs. 3 and 4). The caveat is that these fascinating proposals may apply to some categories of alkaliphile proteins but not to others. Table 3 shows a small selection of alkaliphile exoenzymes and some examples of enzymes other than proteases (e.g., pectate lyase Pel-7; Kobayashi et al., 1999) that have a highly alkaline pH optimum as well as a high pI. However, other enzymes, including other proteases, have high pH optima but not high pIs. Perhaps diverse sets of adaptations are associated with different catalytic activities and mechanisms, different 3-dimensional structural features, or both. Also important is the conjoining, in many alkaliphiles, of the constraints of thermophily or halophily or both with alkaliphily. Thus, the mature form of the a-amylase from haloalkaliphilic Natronococcus sp. Ah-36 was deduced to have a very high acidic amino acid content, but this is a general adaptation to halophily (Lanyi, 1974; Eisenberg et al., 1992).

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25 50 75 100 (S1)

(S2)
(S2)
-AQSVPWGISRVOAPAAHNRGLTGSGVKVAVLDTGI-STHPDLNIR----------------GGASFVPGEPST-QQGNGHGTHVAGTLAALN-NSIGVLGVAPSAELYAVKVLGASGSG -AQSVPWGISRVQAPAAHNRGLTGSGVKVAVLDTGI-STHPDLNIR----------------GGASFVPGEPST-QDGNGHGTHVAGTIAALN-NSIGVLGVAPNAELYAVKVLGASGSG --QTVPWGINPVQAPIAQSRGFTGTGVRVAVLDTGI-SNHADLRIR----------------GGASFVPGEPNI-SDGNGHGTQVAGTTAALN-NSIGVLGVAPNVDLYGVKVLGASGSG i t h t d -QsVPWGISRVQAPAAqSRGFTGTGVKVAVLDTGI-SNHPDLKIR----------------GGASFVPGEPSp-QDGNGHGTHVAGTIAALN-NSIGVLGVAPNAeLYAVKVLGASGSG ancestor A @%@ @ # @ %# AQSVPWGISQIqAPKAqArGYTGTGVKVAViDTGIESNHPDLKiR----------------GGASFVpGEpSPYQDQNGHGTHVAGTIAALN-NSiGVLGVAPSAkLYAVKVLGASGSG ancestor B t k h q i v a a t s d l e B.peptidase F ATDGVENNVDQIDAPKAWALGYDGTGTVVASIDIGVEWNHPALKEKYRGYNPENPNEPENEMNWYDAVAGEASPY-DDLAHGTHVTGTMVGSEPDGTNQIGVAPGAKWIAVKAFSEDG-G sub. Carlsberg -AQTVPYGIPLIKADKVQAQGFKGANVKVAVLDTGIQASHPDLNVV----------------GGASFVAGE-AYNTDGNGHGTHVAGTVAALD-NTTGVLGVAPSVSLYAVKVLNSSGSG -AQSVPYGISQIKAPALHSQGYTGSNVKVAVIDSGIDSSHPDLNVR----------------GGASFVPSSTNPYQDGSSHGTHVAGTIAALN-NSIGVAGVSPSASLYAVKVLDSTGSG sub. J -AQSVPYGVSQIKAPALHSQGYTGSNVKVAVIDSGIDSSHPOLKVA----------------GGASMVPSETNPFQDNNSHGTHVAGTVAALN-NSIGVLGVAPSASLYAVKVLGADGSG sub. BPN’

M-protease PB92 elastase YaB

175 200 125


(S3) (S4) SVSSIAQGLEMAGNNGM-----------HVANLSLGSPSPSA-TLEQAVNSATSRGVLVVAASGNSGA------GSISYPARYANAMAVGATDQNNNRASFSQYGAG------LDIVAPG SVSSIAQGLEMAGNNGM-----------HVANLSLGSPSPSA-TLEQAVNSATSRGVLVVAASGNSGA------GSISYPARYANAMAVGATDQNNNRASFSQYGAG------LDIVAPG SISGIAQGLQWAANNGM-----------HIANMSLGSSAGSA-TMEQAVNQATASGVLVVAASGNSGA------GNVGFPARYANAMAVGATDQMNNRATFSQYGAG------LDIVAPG i g M q r SvSGIAQGLEWAaNNGM-----------HVANMSLGSPSGSA-TTEQAVNsATSsGVLVVAASGNSGA------GSIGYPARYANAMAVGATDQNNNRASFSQYGAG------LDIVAPG ancestor A • %@ % • @ % • % @ % % • • #### @ •@*% • •• @ % TdSGIaqGlEWALANGM-----------DVaNMSLGGPSGSA-tleQAVNAAtSsGVlVVAAAGNSGa--sGGPGSIGYPArYPnamaVGATDsNNNRASFSqyGAg------LdIvAPG ancestor B y le i i amk r r v s p n ssi i lv e e m g v w r f l k evf q sq s s B.peptidase G TDADILEAGEWALAPKDAEGNPHPEMAPDVVNNSWGGGSGLDEWYKDMVNAWRSADIFPEFSAGNTDLFIPGGPGSIANPANYPESFATGATDINKKLADFSLQGPSPYDEIKPEISAPG sub. Carlsberg TYSGIVSGIEWATTNGM-----------DVINMSLGGPSGST-AMKQAVDNYYARGVVVVAAAGNSGS--SGNTNTIGYPAKYDSVIAVGAVDSNSNRASFSSVGAE------LEVMAPG QYSWIINGIEWAISNNM-----------DVINMSLGGPSGST-ALKTVVDKAVSSGIVVAAAAGNEGS--SGSSSTVGYPAKYPSTIAVGAVNSSNQRASFSSAGSE------LDVMAPG sub. J QYSWIINGIEWATANNM-----------DVINMSLGGPSGSA-ALKAAVDKAVASGVVVVAAAGNEGT--SGSSSTVGYPGKYPSVIAVGAVDSSNQRASFSSVGPE------LDVMAPG Sub. BPN’ M-protease PB92 elastase YaB

225 250 275
(S5)
(S6)
VNVQSTYPGSTYA-SLNGTSMATPHVAGVAALVKQKNPSWSNVQIRNHLKNTATGLGNT-------NLYGSGLVNAEAATB--VNVQSTYPGSTYA-SLNGTSMATPHVAGAAALVKQKNPSWSNVQIRNHLKNTATSLGST-------NLYGSGLVNAEAATR--VGVQSTVPGNGYA-SFNGTSMATPHVAGVAALVKQKNPSWSNVQIRNHLKNTATNLGNT-------TQFGSGLVNAEAATR--l VNVQSTVPGSTYA-SLNGTSMATPHVAGVAALVKQKNPSWSNVQIRNHLKNTATNLGNT-------NqYGSGLVNAEAATR--ancestor A • % @ % • •• • @ %@ VNvQSTVPGSTYA-sLNGTSMATPHVAGVAALvKQKNPSwSNvQiRNhLKSTATpLGnS-------NyYGsGLVNAEAAtr--ancestor B i g i l d v r t y d g h aq l m i k vs B.peptidase F VNIRSSVPGQTYEDGWDGTSMAGPHVSAVAALLKQANASLSVDEMEDILTSTAEPLTDSTFPDSPNNGYGHGLVNAFDAVS--sub. Carlsberg AGVYSTYPTSTYA-TLNGTSMASPHVAGAAALILSKHPNLSASQVRNRLSSTATYLGSS-------FYYGKGLINVEAAAG--VSIQSTLPGGTYG-AYNGTSMATPHVAGAAALILSKHPTWTNAQVRDRLESTATYLGNS-------FYYGKGLINVQAAAQ--sub. J VSIQSTLPGNKYG-AYNGTSMASPHVAGAAALILSKHPNWTNTQVRSSLENTTTKLGDS-------FYYGKGLINVQAAAQ--Sub. BPN’ M-protease PB92 elastase YaB

Fig. 4. An alignment of amino acid sequences of M-protease and related proteases showing the sequence of hypothetical ancestors A and B, with the “high-alkaline” protease sequences from Fig. 3 shown above ancestor A and the “alkaline” protease sequences from Fig. 3 shown below ancestor B. Residue numbers of M-protease are indicated. Symbols between the ancestral sequences indicate residues that have been substituted between them. Symbols @, %, and * indicate substitutions of ionizable residues, residues at the interface of shifted segments and others, respectively. Symbol # indicates inserted/deleted residue(s). In the M-protease sequence underlining is used to show positions of shifted sequences in the structural studies. This figure was reproduced, with permission from the publisher, from Shirai et al. (1997).

Table 3. The isoelectric point (pI) and pH optimum for activity of selected exoenzymes from alkaliphilic Bacillus strains. Enzyme Protease PB92 Elastase YaB M-protease ALPase II protease1 Protease Pectate Lyase Pel-7 Xylanase J 1

Bacillus strain

pI

pH optimum

References

B. alcalophilus Bacillus YaB Bacillus KSM-K16 Bacillus NKS-21 Bacillus NKS-21 Bacillus KSMP7 Bacillus sp. str. 41M-1

>10 >10 10.6 2.8 8.2 10.5 5.3

10.5–12 11.7 12.3 10.2 10–11 10.5 9

Vander Laan et al., 1991; Zuidweg et al., 1972 Tsai et al., 1983; Kaneko et al., 1989 Kobayashi et al., 1995 Yamagata and Ichishima, 1989 Tsuchida et al., 1986 Kobayashi et al., 1999 Nakamura et al., 1993

ALP = alkaline proteinase

Are There Global Adaptations of the Cell Surface Layers? Cytoplasmic Membrane Little attention has been focused on the cytoplasmic membrane characteristics of extreme, nonhaloalkaliphilic prokaryotes living in alkaline medium—their primary stress. Indications are that studies of membrane lipids will be important in elucidating major adaptations to such environ-

ments. Koga et al., 1982 and Nishihara et al., 1982 reported on the lipid composition of alkaliphilic Bacillus sp. strain A-007 and several other alkaliphilic Bacillus strains, in which they characterized a novel polar lipid identified as bis-(monoacylglycero)-phosphate (BMP). This lipid’s presence in mammalian lysosomes but not in prokaryotes has been reported (Horikoshi, 1991). Other phospholipids found in Bacillus species included phosphatidylglycerol (PG), phosphatidylethanolamine (PE), and often car-

CHAPTER 1.10

diolipin (CL) with an array of branched-chain and other fatty acids. Cardiolipin concentrations were generally significant, whereas glycolipids and glycophospholipids that often are found in Gram-positive bacteria were absent. The neutral lipid fraction contained diacylglycerol, both squalene and dehydrosqualene, as well as an uncharacterized component that accounted for about 20% of the neutral lipid (Koga et al., 1982). Subsequently, Clejan et al., 1986 compared the lipid composition of several obligately and facultatively alkaliphilic strains. Again, high levels of CL were reported, i.e., 13% and 25% of the polar lipids in B. pseudofirmus RAB and OF4, respectively. The obligately alkaliphilic B. pseudofirmus RAB had a much higher neutral/polar lipid ratio than the facultatively alkaliphilic B. pseudofirmus OF4; the neutral lipid fraction contained squalene and dehydrosqualene, diacylglycerol and some incompletely characterized long-chain isoprenoid lipids. In a subsequent study (Clejan et al., 1988), the permeability of lipid vesicles prepared with different ratios of the diacylglycerol and isoprenoid fractions was found to be enhanced by diacylglycerol and decreased by the isoprenoid fraction. These neutral lipid components may be part of the balancing of fluidity with barrier functions of the coupling membrane. The obligately alkaliphilic B. pseudofirmus RAB also had 90% branched-chain fatty acids as opposed to 72% in the facultatively alkaliphilic B. pseudofirmus OF4, and whereas the facultative alkaliphile had no unsaturated fatty acids in its phospholipids, the obligate alkaliphile had a significant amount (Clejan et al., 1986). It was hypothesized that the obligate alkaliphile might have membrane lipid properties that functioned well at highly alkaline pH values but became too fluid and permeable at near neutral pH values. This hypothesis was supported by the finding that the presence of low concentrations of unsaturated fatty acid stops the facultative strain from growing in the low end of its former pH range for growth and upon incorporation of the unsaturated fatty acid into the membrane (Dunkley et al., 1991). Interestingly, a more recent alkaliphile isolate, Bacillus cohnii was noted to have an extraordinarily high content of unsaturated fatty acids (Spanka and Fritze, 1993). Aono et al., 1992 have noted the instability of protoplast membranes of B. halodurans C-125 at alkaline pH values that are well below the optimum for growth. Similarly, Krulwich et al., 1985a found during studies of cytoplasmic buffering capacities in Bacillus species with diverse pH optima for growth, that the alkaliphiles lost stability below pH 7. These phenomena may reflect the same general membrane property that limits the low range of growth pH. As indicated below,

Alkaliphilic Prokaryotes

291

increased autolysis of the peptidoglycan also could be involved. Recently, Gilmour et al. (manuscript submitted) conducted a study of the proteins found in pH 7.5- and pH 10.5-grown B. pseudofirmus OF4. Among the proteins that were strongly up-regulated at pH 10.5 were two enzymes that are likely to be involved in branched-chain fatty acid metabolism or production. The upregulation of these enzymes might relate to remodeling involved in adaptation to the high pH. The membrane lipids of alkaliphilic Bacillus species clearly contain novel components whose role is unclear, high levels of CL and of branched-chain fatty acids. The more detailed characterization of the membrane lipid components and mutational analyses of their functions will be important areas for furthering our understanding of the physiology of this group.

Peptidoglycan and Associated Polymers The alkaliphile peptidoglycan and associated polymers, especially in Bacillus species, have received more detailed attention than the membrane lipids. The data that have emerged from studies thus far indicate that both the turnover dynamics of the peptidoglycan itself and pHdependent changes in associated, negatively charged polymers may be important contributors to alkaliphile physiology. Koyama and Nosoh, 1976 noted that cells of an alkaliphilic Bacillus strain were more negatively charged upon growth at pH 10 than at 8.2. Subsequent studies of the composition of a diverse group of alkaliphilic Bacillus species at different pH values for growth indicated that these organisms generally have the A1g type of peptidoglycan in which meso-diaminopimelic acid, the third residue in the tetrapeptide of mature peptidoglycan units, is linked via its e-amino group directly to the terminal D-alanine of another peptide (Aono et al., 1984); some variations have since been found, including the presence of ornithine instead of diaminopimelic acid in B. cohnii (Spanka and Fritze, 1993). Some alkaliphiles (e.g. B. halodurans C-125) are more susceptible to autolysis at near neutral pH and exhibit a lower (%) crosslinking of the peptidoglycan at lower growth pH than at alkaline pH (Aono and Sanada, 1994). These interesting differences, however, still do not account for the more negative charge of the surface layers at high pH. Cell wall-associated polymers that are highly acidic appear to be responsible for pHdependent changes in cell surface charge, and the polymers differ between groups of alkaliphilic Bacillus species. Bacillus halodurans C-125 was among those species that produced a teichuronic acid composed of N-acetyl-D-fucosamine, glucu-

292

T.A. Krulwich

ronic acid and galacturonic acid (Aono, 1985) and a teichuronopeptide composed of a polyglucuronic acid and a polyglutamate polymer (Aono, 1989; Aono et al., 1993). Mutants that were defective in one or both of these polymers were growth defective at high pH (Aono and Ohtani, 1990; Ito et al., 1994), which indicates that these polymers are important in the alkaliphily of this group of Bacillus species (Aono and Ohtani, 1990; Aono et al., 1995). Notably, these organisms were cultivated in glucosecontaining media where alkaliphiles are often a bit less fastidious than when in a medium of nonfermentative carbon sources (Gilmour and Krulwich, 1997). The uronic acid polymers are not found in other groups of alkaliphilic Bacillus species, e.g., B. pseudofirmus OF4 (Aono, 1985; Guffanti and Krulwich, 1994). On the other hand, recent studies have shown that B. pseudofirmus OF4 has an acidic S-layer polymer produced from a gene with strong homology to similar genes from Bacillus anthracis and Bacillus licheniformis. This S-layer polymer was identified in 2-dimensional gel electrophoretic patterns of membrane-associated proteins from pH 10.5and pH 7.5-grown cells of B. pseudofirmus OF4, and characterized as a heterogeneous, apparently processed, protein that is present in greater amounts at high pH. By cloning, sequencing and disrupting the S-layer gene, the S-layer is found to be dispensable for alkaliphily in B. pseudofirmus OF4, but confers an advantage to cell growth at pH 10.5 and to cytoplasmic pH homeostasis in a sudden shift from pH 8.5 to 10.5 (Gilmour et al., manuscript submitted). B. pseudofirmus OF4 also may have an acidic capsule layer because a partial sequence for genes that are likely to encode enzymes synthetic for a polyglutamate capsule was identified in this species (Ito et al., 1997a). The characterization of the complete sequence and role of this locus will be interesting.

Are There Global Adaptations of Cytoplasmic Components? Buffering Capacity A comparison of the cytoplasmic buffering capacity of Bacillus species that grow in vastly different pH ranges indicated that alkaliphiles, grown on nonfermentable carbon sources, had higher cytoplasmic buffering capacities at alkaline pH than at lower pH (Krulwich et al., 1985a). In a recent study, Rius and Loren, 1998 reported comparative values for the cytoplasmic buffering capacity of B. alcalophilus grown on both fermentative and nonfermentative carbon

CHAPTER 1.10

sources. These investigators used the decay of an acid pulse (Maloney, 1979) to determine both cytoplasmic buffering capacity and membrane H+ conductance, thereby avoiding problems associated with permeabilizing cells. In media with either nonfermentative carbon sources or fermentative carbon sources, the cytoplasmic buffering capacity of B. alcalophilus was much higher in pH 10.5-grown than in pH 8.5-grown cells. Strikingly, the alkaliphile cells (and others, such as Staphylococcus aureus and B. subtilis) had vastly lower cytoplasmic buffering capacity when grown on malate-carbonate media than on media with fermentative carbon sources (Rius and Loren, 1998). Since malate-carbonate media support a more alkaline optimum for the growth pH, there is probably no direct relationship between overall cytoplasmic buffering capacity and the capacity to grow at the upper reaches of pH. On the other hand, it is quite possible that specific compounds play particular roles in connection with alkaliphily. For example, several studies have focused on a shift in the ratio of the major polyamine compounds such as spermidine, which predominates heavily at very alkaline pH values (Chen and Cheng, 1988; Hamana et al., 1989).

Alkali-Stability of Cytoplasmic Components The general impression is that the cytoplasmic pH, for most alkaliphiles in their optimum pH range, is within about 0.5 pH units of the cytoplasmic pH optimum for most bacteria. Hence there may not be major or even discernible, global adaptations in the protein structure or pH profile (Horikoshi and Akiba, 1982; Horikoshi, 1991). But throughout the literature on alkaliphiles, at least some apparent cytoplasmic enzymes have unusually high pH profiles (Horikoshi and Akiba, 1982). Some reports would seem to beg for an explanation; for example, a putative “intracellular alkaline serine protease” from alkaliphilic Thermoactinomyces sp. HS682 was produced from its gene in Escherichia coli. The purified enzyme had a pH optimum of 11 (Tsuchiya et al., 1997). In consideration of such observations and the recognition that some of the soda lake alkaliphiles have evolved over a long period in a consistently high alkaline environment, it may be well worth looking explicitly for exceptions to the expectation that cytoplasmic proteins have no global adaptations to a higher than conventional pH. In a non-alkaliphile that completely lacks active pH homeostatic mechanisms, such as Clostridium fervidus (Speelmans et al., 1993), the organism can only grow in a narrow pH range up to about pH 7.7. In the

CHAPTER 1.10

especially effective pH homeostatic mechanisms of most well studied alkaliphiles (Krulwich et al., 1997), the upper pH limit is generally about pH 9 to 9.5. Even in those alkaliphiles, which have most often been soil isolates originally, subtle but important global adaptations in the cytoplasmic proteins are possible. Perhaps among the soda lake alkaliphiles there are organisms that are less dependent on remarkable pH homeostasis mechanisms because their cytoplasmic enzymes and functional assemblies (e.g., secretion and protein synthetic machinery) are all markedly alkali-adapted relative to those of conventional bacteria. A novel osmolyte has been noted in haloalkaliphilic archaea (Desmarais et al., 1997). Perhaps the cytoplasmic proteins of some of these organisms are not only salt-adapted but also at least unusually alkali-adapted. It also will be of interest to carefully examine the properties of ribosomal and other proteins that must interact with nucleic acid molecules in a cytoplasm that is generally at least half a pH unit higher than the cytoplasm of conventional bacteria growing at optimal pH (Krulwich, 1995). Horikoshi, 1991 has noted a pH optimum for protein synthesis in an alkaliphilic Bacillus was about half a pH unit higher than that of B. subtilis. Genes for ribosomal proteins and major polymerases are beginning to be identified so that detailed examinations will be facilitated (Nakasone et al., 1998; Takami et al., 1999d). Thus far, homologous DNA-binding proteins from alkaliphiles and non-alkaliphiles have not been compared in great detail, the two small acid-soluble spore proteins (SASPs) sequenced from alkaliphiles (Quirk, 1993; Wei et al., 1999) have not been examined closely from this perspective.

Proteome Studies The 2-dimensional gel analysis of proteins from steady-state pH 7.5- and pH 10.5-grown B. pseudofirmus OF4 cells, and from cells grown at lower pH and rapidly shifted to pH 10.5, revealed several interesting features that will merit further examination in this and other alkaliphiles (Gilmour et al., manuscript submitted). Significant numbers of proteins were found to be either up-regulated or down-regulated at high pH, but the extent of the change was typically greater for the up-regulated genes. In addition, a substantially greater number of genes were upregulated transiently with rapid increase of pH to 10.5 than were ultimately up-regulated in the steady-state cells; this find correlates with other data indicating that there are groups of proteins that play a specific role in the initial adjustment to a major alkaline shift.

Alkaliphilic Prokaryotes

293

Genomics The vastly increasing database on sequences of alkaliphile proteins will help clarify the issues already raised about global adaptations in cytoplasmic proteins or in functional cytoplasmic assemblies (e.g., ribosomes and secretory particles). The completion of the B. halodurans C-125 genome will be a major step in this process (Takami et al., 1999b, Takami et al., 1999d). The maps of alkaliphiles and gene order in several large pieces of DNA suggest that interesting features of alkaliphily may emerge (Takami et al., 1999b; Takami et al., 1999d; Gronstad et al., 1998; Wei et al., 1999). Some, but probably not the majority, of the alkaliphilic Bacillus species and strains have been found to harbor endogenous plasmids. Most of these have been relatively small and have not been exciting candidates for development of host-vector systems. Rather, the plasmids that have been extensively developed for use in B. subtilis generally have been adapted and applied to molecular manipulations of alkaliphilic Bacillus species (Horikoshi, 1991). However, Fish et al., 1999 have recently found that eight alkaliphile halomonads, out of the seventeen they examined, possessed one or more plasmids in the size range of 5.3 to 33 kb; they concluded that some of these would be suitable for vector development. A large endogenous plasmid, approximately 30 kb, also has been found in B. pseudofirmus OF4 (Gronstad et al., 1998) and bears the cadmium-resistance locus (Ivey et al., 1992). There is no evidence to date for a role of plasmid-borne genes in alkaliphily.

Active pH Homeostasis and the Involvement of Secondary Na+/H+ Antiporters and Secondary Na+/Solute Symporters In alkaliphilic Bacillus species, active ion transport mechanisms are central to the crucial process of pH homeostasis, and this process appears to limit the upper pH limit for growth (Krulwich, 1995; Krulwich et al., 1997). Sturr et al., 1994 conducted studies of the bioenergetic parameters of B. pseudofirmus OF4 in a pH-controlled continuous culture apparatus in which the bacteria were grown aerobically on malate-containing media at various, fixed pH values from pH 7.5 to 11.4. In fact, the upper limit for growth was not found in this range, with a long but certainly viable generation time of 700 min found at pH 11.4. As shown in Fig. 5, B. pseudofirmus OF4 grows slightly better at pH 8.5 to 10.5 than at pH 7.5, and throughout this pH range, the cytoplas-

294

T.A. Krulwich

CHAPTER 1.10

Problem: Regulation of Cytoplasmic pH at Very Alkaline Values of Growth pH is Extraordinarily Effective, and May Become Limiting to Growth Only Above pH 11

Problem: Oxidative Phosphorylation at High pH Values at which the Chemiosmotic Driving Force is Low Works Even Better than at Lower pH Values at which the Putative Driving Force is About Three Times Higher

700

∆ψ or ∆p, mV –180

600

–160

tg. min Cytoplasmic pH

10

∆ψ (1.8)

500

–140 (13)

9

400

(2.3)

300

–100 (3.2)

(2.3) (∆pH)

8

–120

200

(0.8) (0)

0 7.5

(7.2)

∆Gp

pHin

–500 –400

–80 ∆p

(2.0)

100 7

(∆Gp/∆p) (7.9)

(11)

∆Gpt mV –600

–60

8.5 9.5 10.5 pH of Medium

11.5

–40 7.5

8.5 9.5 10.5 pH of Medium

mic pH remains below 8.5; at external pH values of 9.5 and above, the dpH (transmembrane pH gradient) is 2 full pH units or more. At the highest external pH included in the study, pH 11.4, the internal pH was at about 9.6. The growth rate slowed dramatically, and in parallel, with the increasing cytoplasmic pH above a value of 8.2. Thus in this type of alkaliphilic Bacillus, pH homeostasis is remarkable: cytoplasmic pH is maintained at about pH 7.5 until the dpH exceeds 2 and thereafter, the rise in cytoplasmic pH is correlated with a decrease in growth rate (i.e., increase in generation time). The finding of optimal growth rates up to about pH 10.5, at which the cytoplasmic pH is maintained below pH 8.5, was corroborated in other chemostat studies (Guffanti and Hicks, 1991) as well as batch culture studies (Hirota and Imae, 1983; Aono et al., 1997). Under conditions in which the cells are well energized, the magnitude of the dpH found among different alkaliphilic Bacillus species using several different kinds of probes to assess this parameter has been very consistent. Significantly, the full dpH that the alkaliphiles maintain was not measured in the earliest studies (e.g., Guffanti et al., 1978) because the measurements were carried out in buffers without energy sources. Thus compendia of such measurements often contain citations to a mixture of experimental conditions which determine the extent to which alkaliphiles grow. There is a large body of evidence for the crucial Na+ cycle involvement in net acidification of the cytoplasm during growth of alkaliphilic prokaryotes at alkaline pH. That alkaliphiles required Na+ and used Na+ as the coupling ion for transport systems (Koyama et al., 1976;

11.5

Fig. 5. Bioenergetic parameters of B. pseudofirmus OF4 during growth in continuous cultures at various controlled pH values. The data from Sturr et al., 1994 were replotted to highlight particular features of interest. Left: the doubling time (tg, min) and cytoplasmic pH (pHin) are shown as a function of the growth pH. The numbers in parentheses above the points on the cytoplasmic pH curve are the values for dpH. Right: The transmembrane electrical potential, positive out (d y), the total bulk electrochemical proton gradient (dp), and the phosphorylation potential (dgp) are all shown in mV as a function of growth pH. The numbers in parentheses are the dGp/dp, which would reflect the H+ stoichiometry if coupling were strictly to a bulk gradient. This figure was reproduced with permission from the publisher from Krulwich (1995).

Kitada and Horikoshi, 1977; Guffanti et al., 1978) had already been demonstrated when work on both wild type alkaliphiles and pH homeostasis-negative non-alkaliphilic mutants began to implicate Na+/H+ antiporters as key mediators of cytoplasmic acidification (Mandel et al., 1980; Koyama et al., 1986; Krulwich and Guffanti, 1983, Krulwich and Guffanti, 1989). That maintenance of a cytoplasmic pH well below the external pH depends upon the availability of Na+ (Mandel et al., 1980; Krulwich et al., 1982; McLaggan et al., 1984; Krulwich et al., 1985b) was established in several types of alkaliphiles. Diverse mutant strains, non-alkaliphiles that were specifically deficient in their ability to regulate cytoplasmic pH, also were shown to be deficient in Na+/H+ antiport (Mandel et al., 1980; Krulwich et al., 1982; Garcia et al., 1983; Hamamoto et al., 1994; Krulwich et al., 1996). Conversely, B. pseudofirmus OF4 cells taken from continuous cultures maintained at pH 11.4, contained variants with elevated Na+/H+ antiporter activity (Sturr et al., 1994). Electrogenic Na+/H+ antiporters that catalyze exchange of intracellular Na+ for a stoichiometrically greater number of H+ from the external milieu transport net positive charge inward during each turnover. Accordingly, these fluxes can be energized by the transmembrane electrical potential component (d) of the total electrochemical proton gradient (dp) that is established by the proton-extruding respiratory chain (or by H+-extruding ATPases in anaerobic cells). Components of the respiratory chain and diverse, electrogenic Na+/H+ antiporters are depicted in a diagram of an aerobic alkaliphilic Bacillus in Fig. 6. Because the d can energize the electro-

CHAPTER 1.10

Alkaliphilic Prokaryotes

295

C-125 Teichuronic acid Teichuronopeptide xH+ bd MQ

II

I, nuo

T.O. Respiration

NatB A

ATP ADP + pi

NatB C

ATP ADP + pi

Na+

?

T.O.

ATP synthase ATP

F1

Solute C

Alkaliphile membrane lipid adaptations

S

F0

caa3

C

Na+

Na+ S Solute D

Q: Are the cytoplasmic enzymes and protein assemblies alkali-adapted in the most highly adapted and specialized soda lake alkaliphiles? Q: Are there special osmolytes, buffers?

Electrogenic

mot

Na+

Na+/H+ antiporters

CF4

MrpA

NapA

NhaE

S

S

nH+ qNa+

rH+ jNa+

kH+ Na+ Solute Na+ Solute B A

+

mNa+

a

N

S-layer capsule

ot

m



xH+

ADP + pi

NhaC S

MQ III

?

xNa+ yH+

xH+

Peptidoglycan

Fig. 6. A schematic of an alkaliphilic Bacillus, showing the membrane-associated ion-translocating proteins and complexes involved in the primary generation of a dp as well as the secondary antiporters (Mrp, Nap, Nha) and symporters (S) that together catalyze solute uptake and net proton accumulation to achieve a lower cytoplasmic than medium pH. Also shown are: the flagellar assembly and associated Na+ channel that may provide another physiologically important Na+ reentry route in addition to the Na+-coupled symporters; and the F1FO-ATP synthase. This proton-coupled synthase may, at pH > 9.2, accept protons that are somehow sequestered as indicated by arrows parallel to, and either just above or just below, the membrane surface. The dotted outline of the caa3-type oxidase represents a hypothetical route by which protons are transferred directly from this complex to the synthase in protein-protein interactions. The dotted outline of MrpA indicates that at least under some conditions this antiporter may function as part of a complex. Outlines of a cell-wall-associated layer reflect the finding that in B. halodurans C-125 and B. pseudofirmus OF4, different negatively charged polymers play at least some role in pH homeostasis. Other possibilities that are yet to be clarified are indicated by the questions (Q:) presented in the cytoplasmic space.

genic antiport, it is possible for such antiporters, working in concert with respiration, to acidify the cytoplasm relative to the medium (McNab and Castle, 1987). For such antiport to support pH homeostasis on a continuous basis, however, Na+ must be recycled to maintain the source of the cytoplasmic substrate for the Na+/H+ antiporters. Compelling evidence has been presented for an important role of the numerous Na+-coupled solute uptake systems, Na+/solute symporters, in Na+ reentry in support of pH homeostasis (Krulwich et al., 1985b). In fact, it was hypothesized, when pleiotropy was observed in some non-alkaliphilic mutant strains, that the alkaliphile Na+/H+ antiporter(s) and Na+/solute symporter(s) playing critical roles in pH homeostasis might share a common subunit (Guffanti et al., 1981). However, both the ion specificity of the transporters (Sugiyama et al., 1985) and the many genes that now have been identified fail to support such a common structure. Rather, the

pleiotropy of many non-alkaliphilic mutants likely relates to the complexity of the physiological networks surrounding the important cell functions of pH homeostasis, solute transport and Na+-resistance (Krulwich et al., 1996, Krulwich et al., 1997, Krulwich et al., 1998) and the likely complexity of one of the major antipor/t systems (Hamamoto et al., 1994; Hashimoto et al., 1994; Ito et al., 1999). In addition to the symporters, some pH-sensitive mechanism for Na+ reentry must complete the Na+ cycle that supports pH homeostasis even when solutes are not present and Na+ is not abundant (Booth, 1985; Krulwich and Guffanti, 1989; Krulwich et al., 1997). A good candidate for a pH-dependent Na+ entry route of this sort, as depicted in Fig. 6, is the Na+-translocating channel associated with flagellar rotation in the alkaliphilic Bacillus species (Sugiyama, 1995). Although much less complete pictures of the pH homeostasis cycle of alkaliphiles outside the genus Bacillus have been

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presented, parameters of the cycle (Cook et al., 1996) and involvement of Na+ or Na+/H+ antiporters or both in the central cell energetics of other alkaliphiles have been described (Buck and Smith, 1995; Prowe et al., 1996). The cycle outlined above for pH homeostasis has been the model of this function in many nonalkaliphilic bacteria, but the alkaliphilic Bacillus species have an extraordinary capacity for pH homeostasis as well as some specificity that is not shared by non-alkaliphiles. B. pseudofirmus OF4 has at least 10 times the aggregate Na+/H+ antiporter activity as B. subtilis and has far more capacity for pH homeostasis than the non-alkaliphile. In addition, pH homeostasis in B. subtilis can draw upon K+/H+ antiport as well as well as Na+/H+ antiport, which is carried out by antiporters that can use either cation. On the other hand in the alkaliphilic Bacillus species, the process depends specifically on Na+ (Krulwich et al., 1994, Krulwich et al., 1999). Although there is evidence for K+/H+ antiporters in alkaliphiles (Mandel et al., 1980; Kitada et al., 1997), and the first Na+(K+)/H+ antiporter-encoding gene has now been identified in B. pseudofirmus OF4 (Wei et al., manuscript submitted), it is unclear what the role of the K+/H+ antiport is and why it does not come into play in any obvious way in pH homeostasis. If B. subtilis is subjected to a sudden shift in pH from 7.5 to 8.5, for example, the initial cytoplasmic pH of about 7.5 is maintained as long as either K+ or Na+ is present. Whereas in the alkaliphile, only Na+ can support this homeostasis or comparable homeostasis during a pH 8.5 to 10.5 shift in the external pH (Krulwich et al., 1994; Krulwich et al., 1999; Ivey et al., 1998). Most work on pH homeostasis of alkaliphilic Bacillus species has lately been carried out on B. halodurans C-125 and B. pseudofirmus OF4 and has focused upon identification of the antiporter-encoding genes that have roles in pH homeostasis. It has been clear for several years that, as in other bacteria (Padan and Schuldiner, 1996; Krulwich et al., 1999), alkaliphilic Bacillus species have more than one Na+/H+ antiporter system with a role in pH homeostasis (Kitada et al., 1994; Krulwich, 1995; Ito et al., 1997b). Currently, more Na+/H+ antiporters have been identified in B. pseudofirmus OF4 than in any other single organism; these are indicated in Fig. 6 as mrp, nhaC, napA, and nhaE. It is likely that one of these antiporters has the dominant role in pH homeostasis. This antiporter is encoded by a locus first identified and partially characterized by Horikoshi, Kudo and colleagues in B. halodurans C-125 (Kudo et al., 1990; Hamamoto et al., 1994). It is also found in B. pseudofirmus OF4 (Krulwich et al., 1998; Krulwich et al., 1999) and other bacteria

CHAPTER 1.10

(Putnoky et al., 1998; Hiramatsu et al., 1998) and has been named mrp (for multiple resistances and pH) in B. subtilis (Ito et al., 1999). The locus encoding all of these homologues is an intriguing 7-gene operon in which all of the deduced products are hydrophobic and several of them have sequence similarity to membraneembedded subunits of the NDH-1 type of NADH dehydrogenase complexes (Yagi, 1993). A point mutation in the mrpA gene of B. halodurans C-125 renders the bacteria nonalkaliphilic and extremely defective in pH homeostasis (Hamamoto et al., 1994). Deletion of the mrpA of B. subtilis makes this strain extremely sensitive to Na+ and modestly deficient in pH homeostasis in certain concentration ranges of Na+ or K+ (Ito et al., 1999). An unusual dependence of MrpA function on other genes in the operon (Hiramatsu et al., 1998; Ito et al. 1999) has led to the speculation that mrp might be an obligatory complex (Hiramatsu et al., 1998). Because it appears that MrpA can function as a secondary antiporter system in stoichiometric excess but not in the absence of the other gene products, another complexinvolved mode of transport is still possible (Ito et al., 1999); this is indicated by the dotted lines in Fig. 6. The dissection of the structural form(s) of the active complex, their mechanism and roles in the alkaliphile, and their differences from homologues in non-alkaliphiles, are important areas of current investigation. The nhaC gene of B. pseudofirmus OF4 was the first alkaliphile Na+/H+ antiporter-encoding gene to be cloned, which was achieved by functional complementation of an antiporterdeficient Escherichia coli mutant (Ivey et al., 1991). Subsequent deletion of the gene showed that the antiporter played a role in high affinity Na+/H+ antiport at both pH 7.5 and 10.5. It was the major antiporter with such affinity in pH 7.5grown cells, whereas another high affinity system was induced during growth at pH 10.5 and the constitutive higher affinity antiport (MrpAassociated?) was preserved (Ito et al., 1997a). An antiporter that confers both Na+/H+ and K+/H+ antiporter activities upon E. coli has been designated nhaE; this antiporter also has a modest role at both pH 7.5 and 10.5 and is in fact expressed more at the lower pH as evidenced by northern analyses (Wei et al., manuscript submitted). Finally, a homologue of the napA antiporter, first described in Enterococcus hirae by Waser et al., 1992 and for which a homologue was subsequently reported in Bacillus megaterium (Tani et al., 1996), also has been found in B. pseudofirmus OF4 and shown to restore Na+-resistance to antiporter-deficient E. coli strains (Wei, Y. and Ito, M. unpublished data).

CHAPTER 1.10

One of the key tasks in further clarifying the basis for the alkaliphile’s capacity for pH homeostasis is the enumeration of all the antiporters involved in a single species and the determination of their roles, mechanisms, and interplay. Another area of interest will be the development of information at a similar molecular level about the reentry routes; do secondary Na+/solute symporters that play a role in pH homeostasis as well as simple solute uptake have particular properties associated with this dual role? Also, it will be important to further examine aerobes growing on fermentative substrates, where some sparing of Na+-dependent, respiration-driven pH homeostasis has been observed (Gilmour and Krulwich, 1997) and to characterize the process in a broader spectrum of extreme alkaliphiles.

Primary Membrane Transport and Motility Primary Active Membrane Transport Systems Secondary active transport systems are centrally important both to Na+-coupled uptake of many solutes and to a Na+ cycle that functions in pH homeostasis. Increasingly, primary active transport systems also are being characterized in diverse alkaliphiles. Among the ATP-driven systems are: Na+-translocating ATPases, at least one of which is a V-type system (Koyama, 1996; Kaieda et al., 1998; Prowe et al., 1996); a Na+-translocating ABC-type transport system (natCAB; Wei et al., 1999) that is homologous to a similar system in B. subtilis (Cheng et al., 1997), and P-type ATPases that confer Cd+- (Ivey et al., 1992) and Na+-resistance (Koyoma, 1999). Other apparent ABC-type systems have been noted in sequences already presented, but studies that might indicate whether alkaliphile transporters have any common features have not been reported as yet. Among the haloalkaliphiles, retinal-based, light-driven primary transport systems have been found and characterized. Although the absence of bacteriorhodopsin-like pigments from these organisms was reported (Bivin and Stoeckenius, 1986), the chloride pump, halorhodopsin, has been characterized in considerable detail (Lanyi et al., 1990; Scharf et al., 1994; Varo et al., 1995; Varo et al., 1996).

Motility Aono et al. (1992) noted that B. halodurans C-125 produced flagella only in the more alkaline part of its pH range for growth, and Sturr et al., 1994 made comparable observations on B. pseudofirmus OF4. The hag gene of the former

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alkaliphile has been sequenced (Sakamoto et al., 1992), whereas the amino acid composition of the flagellin from the latter alkaliphile was determined (Guffanti and Eisenstein, 1983). In both instances, a rather low calculated pI was observed relative to homologues from nonalkaliphiles. The identification of the remaining motility and flagellar assembly genes awaits completion of more genomic studies of alkaliphiles, but the mot assemblies were visualized by rapid freeze electron microscopy (Khan et al., 1992). Imae and his colleagues (Hirota et al., 1981; Hirota and Imae, 1983; Sugiyama et al., 1986) first showed that motility in alkaliphilic Bacillus species was energized by an electrochemical gradient of Na+ as opposed to the dp-driven systems of most non-alkaliphiles. Indeed, amiloride and some of its analogues, inhibitors of various Na+ translocation pathways in eukaryotes, were found to inhibit alkaliphile flagellar rotation (Sugiyama et al., 1988; Atsumi et al., 1990). Because, however, the molecular characterization of the motility-related genes in alkalinetolerant marine bacteria that also use an electrochemical Na+ gradient is incomplete, extensive recent progress on specific properties of the Na+-dependent motility mechanism has occurred in those systems rather than in alkaliphiles (McCarter, 1995; Yorimitsu et al., 1999).

Oxidative Phosphorylation Respiratory Chain Although diverse marine bacteria have been shown to have respiration-coupled, primary Na+ extrusion systems (Tokuda and Unemoto, 1984; Tomb et al., 1993; Beattie et al., 1994; Skulachev, 1994; Pfenninger-Li et al., 1996; Park et al., 1996), thus far the non-marine, nonhalophilic alkaliphiles—including the best studied Bacillus species—have been found to have H+-translocating respiratory chains (Lewis et al., 1983; Krulwich and Guffanti, 1989). These respiratory chains are often branched, with multiple terminal oxidases, and the component cytochrome and iron sulfur protein components are characteristically present at high levels in the membranes (Hicks and Krulwich, 1995; Krulwich et al., 1998). The H+-translocating respiratory chains may facilitate the support of pH homeostasis and the extra costs of oxidative phosphorylation at high pH; but, notably, the alkaliphilic Bacillus species have high molar growth yields on malate (Guffanti and Hicks, 1991; Sturr et al., 1994). In addition to the high concentration of membrane cytochromes and other respiratory chain components, especially at high pH (Lewis et al., 1981; Quirk et al., 1993;

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CHAPTER 1.10

Hicks and Krulwich, 1995; Aono et al., 1996), the respiratory chain components of alkaliphiles have characteristically low midpoint potentials (Lewis et al., 1981; Kitada et al., 1983; Yumoto et al., 1991) as shown for c-type cytochromes in Fig. 7. Yumoto et al., 1991 have suggested that the low midpoint potentials may facilitate electron movement in the inward direction in membranes that maintain a rather high (positive out) transmembrane electrical potential. The respiratory chain components that have been characterized in alkaliphilic Bacillus species have been summarized (Hicks and Krulwich, 1995). Although there is some indirect indication a proton-translocating NADH dehydrogenase (complex I) is present (Hicks and Krulwich, 1995), this is far from established and awaits compelling biochemical and genomic evidence one way or the other. On the other hand, succinate dehydrogenases have been purified and characterized (Gilmour and Krulwich, 1996; Qureshi et al., 1996) and an incompletely characterized bc complex is evident (Lewis et al., 1981; Reidel et al., 1993). Also evident are diverse terminal oxidases, including a pHregulated caa3-type oxidase in B. pseudofirmus OF4 (Quirk et al., 1993), a comparable enzyme recently purified from Bacillus YN-1 (Higashibata et al., 1998), a bd-type cytochrome oxidase in the same species (Gilmour and Krulwich, 1997), and an aco3-type oxidase in Bacillus YN2000 (Qureshi et al., 1990; Yumoto et al., 1993). Engelhard and colleagues (Scharf et al., 1997) have begun to dissect the respiratory chain of the haloalkaliphilic archaeon Natronobacte-

Midpoint potential (mV)

0

RAB(m) RAB c-552(p)

+100

Alkaliphiles

YN YN c(aco )(p) 3 c553(p)

sub c-550(p)

+200

alc(m) YN c-552(p)

PS3 c(bf)(p)

PS3 PS3 c-551(p) c(caa3)(p) sub c-554(P)

Neutralophiles

+300 Fig. 7. Midpoint potentials from various alkaliphile ccytochromes [RAB, B. pseudofirmus RAB; alc, B. alcalophilus; YN, Bacillus YN-2000] in comparison to those from non-alkaliphilic Bacillus species [sub, B. subtilis; PS3, Bacillus PS3]; m (composite membrane data); and p (purified protein). This figure was reproduced with permission from the publisher from Hicks and Krulwich, 1995.

rium pharaonis and note the high respiratorychain component content and possible presence of bc and ba3 complexes; these investigations should help establish what may be more general properties of respiratory complexes in diverse alkaliphiles. While the status of energy-transducing NDH-1 type NADH dehydrogenases is still equivocal in alkaliphiles, the status of NDH-2 type NADH dehydrogenases are not (Xu et al., 1989; Niimura et al., 1995; Aono et al., 1996; Koyama et al., 1998). An interesting set of findings and proposals is emerging from recent studies that bring together the issue of peroxide toxicity, catalase, and the alkaliphile respiratory chain. Yumoto et al., 1990 first purified a protoheme-containing catalase from alkaliphilic Bacillus YN-2000 and reported that it was present in higher activity at elevated pH. In B. pseudofirmus OF4, a complicated profile of three different catalase isozymes has been presented by Hicks, 1995 who also showed that this alkaliphile was more sensitive to killing by hydrogen peroxide at pH 10.5 than at pH 7.5, even though the aggregate catalase activity was about two-fold higher at pH 10.5. Niimura et al., 1995 observed that the NADH oxidase from A. xylanus, which, as described above, is cytochrome-deficient, reduces oxygen to hydrogen peroxide. When the 22-kDa AhpC disulfidecontaining protein from Salmonella typhimurium was added to the reactions, the hydrogen peroxide was reduced to water. Under those circumstances, the net reaction is the oxygendependent oxidation of NADH with the production of NAD+ and water. The NADH-2 from Bacillus YN-1 has now been shown to have similar properties to the A. xylanus enzyme, and there is an AhpC candidate upstream (Koyama et al., 1998). Thus NDH-2, as well as catalases, may be part of a detoxification system. Studies of cyanide-sensitivity of oxygen reduction by obligately alkaliphilic Bacillus YN-1 (Higashibata et al., 1998) have indicated that the cyanide-sensitive component that is attributed to the caa3-type terminal oxidase represents only 10% of the total. The majority, a cyanideinsensitive component, was associated with a low-molecular-weight nonproteinaceous material that has not been completely characterized. The investigators propose a model of alkaliphile respiration in which this cyanide-insensitive terminal respiratory component, in concert with catalase, is of major importance in the respiratory mechanisms.

Respiration-Dependent ATP Synthesis Perhaps in the haloalkaliphilic Archaea, with primary light-driven pumps and unusual mem-

CHAPTER 1.10

brane lipids, very high transmembrane electrical potentials are generated to offset the dpH (acid in) that alkaliphiles maintain. In the alkaliphilic aerobic Bacillus species, however, oxidative phosphorylation presents a clear conundrum, which relates to the higher total chemiosmotic driving force for a proton-coupled process, the electrochemical proton gradient or dp, at pH 7.5 than at pH 10.5. Most measurements (Hirota and Imae, 1983; Sturr et al., 1994) indicate that at the lower pH, the dp is about 3-times the magnitude of that at pH 10.5. Yet the phosphorylation potential, proportional to [ATP]/[ADP][Pi], which reflects the ATP sustained at equilibrium, is greater at pH 10.5 than at pH 7.5 (Fig. 5, right). While Hoffmann and Dimroth, 1991b have calculated somewhat lower discrepancies, their data on ATP synthesis by B. alcalophilus nonetheless show clearly better synthesis at higher pH values where the dp is lower than at near neutral pH values. Moreover, unlike motility and solute symport systems, ATP synthesis by alkaliphilic Bacillus species such as B. alcalophilus and B. pseudofirmus OF4 does not “avoid” the problem of the low electrochemical proton gradient, dp, by using a larger electrochemical Na+ gradient instead. The F1FO-ATP synthases of these organisms were purified and functionally reconstituted into proteoliposomes (Hicks and Krulwich, 1990; Hoffmann and Dimroth, 1991) and shown to be H+- and not Na+-coupled. Thus while there do exist Na+-coupled ATP synthases in several anaerobic marine organisms (Kluge et al., 1992; Forster et al., 1995), the alkaliphilic Bacillus species have not elected that solution. Yet they grow at high molar growth yields on non-fermentative substrates. This suggests that ATP synthesis either operates with a variable coupling stoichiometry, or that it operates well out of equilibrium with the bulk dp because some sort of proton sequestration or other basis for disequilibrium exists. The possibility of a variable stoichiometry merits ongoing consideration and has been proposed for H+-coupled synthesis of ATP in cyanobacteria (Krenn et al., 1993; Van Walraven et al., 1997). In the alkaliphilic Bacillus species, though, the conundrum is further delineated by the observation that respiration-dependent ATP synthesis proceeds well at pH values above about 9, only when respiration itself is the energy source (i.e., artificially imposed gradients of the same magnitude are not efficacious; Guffanti et al., 1984; Guffanti and Krulwich, 1992; Guffanti and Krulwich, 1994; Ivey et al., 1998). This would not be expected if a variable stoichiometry could be employed. Moreover, Ivey et al., 1994 presented data that supportthe actuality of one ATP synthase assembly with an invariant subunit stoichiometry in B. pseudofirmus OF4.

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A variety of sequestration models also merit consideration. Although discrete organelles of various sorts were suggested (Skulachev, 1991), fine structural evidence does not support the presence of pronounced organelles (Rhode et al., 1989; Sturr et al., 1994). Alternatives that are schematically suggested in Fig. 6, include some help from trapping of protons by cell-wallassociated polymers; however, the parameters of oxidative phosphorylation are present in rightside-out membrane vesicles that lack peptidoglycan assemblies (Guffanti and Krulwich, 1994) and oxidative phosphorylation is not restricted at least in B. pseudofirmus OF4, mutants lacking the S-layer (Gilmour et al., manuscript submitted). Protons might also be sequestered by being transferred from a proton-pumping respiratory chain complex, such as the caa3-type oxidase shown in Fig. 6, during a direct protein-protein interaction in the near membrane region of the phospholipid headgroups (Krulwich, 1995). Alternatively, protons might move rapidly along the surface as has been observed in some experimental systems (reviewed by Gutman and Nachliel, 1995). Such translocation, however, should be sensitive to ionic strength, which was not the case for oxidative phosphorylation by ADP and phosphate (Pi)-loaded right-side-out vesicles of B. pseudofirmus OF4 (Guffanti and Krulwich, 1994). It is notable that features, referred to as “alkaliphile-specific sequence motifs,” have been found in important membrane-associated regions of FO subunits of several alkaliphilic Bacillus species (Ivey and Krulwich, 1991, Ivey and Krulwich, 1992; Krulwich et al., 1998). Thus one of the interesting experimental approaches that can be used to test various hypotheses in connection with the energization of alkaliphile oxidative phosphorylation is to alter these motifs and determine whether synthesis is particularly affected at highly alkaline pH.

Why Do Alkaliphiles Generally Grow Poorly or Fail to Grow at Near Neutral pH? There are several indications that the alkaliphiles with the very highest upper edge or highest pH optimum for growth also may be obligate alkaliphiles that cannot grow at pH values much below 9 or 9.5. However, this conjecture has not been examined rigorously in carefully pHcontrolled continuous culture conditions. In batch cultures, for example, Dunkley et al., 1991 noted that an obligate alkaliphile “outcompeted” a closely related facultative alkaliphile at pH 10.5. The putative inverse

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relationship between maximally effective alkaliphily and the ability to thrive at conventional pH values can be viewed as adaptations that foster growth at the extreme but are disabling at lower pH values (“inverse adaptiveness”). Alternatively, the adaptation to high pH may not be directly injurious at lower pH values but constitute an energy cost that compromises growth (“irrelevant cost”). Or, growth at near-neutral pH may elicit essentially a “stress response” among those alkaliphiles that are maximally adapted to optimal growth at highly alkaline pH; some of the consequent shutdown, which is then manifest as a poorer growth rate, could be seen as a stress adaptation (“neutral pH as stress”). Possible examples of each of these may be found in connection with the cell surface. An example of the “inverse adaptiveness” scenario would be the hypothesis (Clejan et al., 1986; Clejan et al., 1988; Dunkley et al., 1991) that obligate alkaliphilic Bacillus strains have bulky branchedchain and unsaturated fatty acids that are adaptive for growth at the upper edges of their pH range but which render the membrane unstable at neutral pH. A possible example of the “irrelevant cost” hypothesis is the finding that the deletion of the S-layer gene from B. pseudofirmus OF4 causes a modest growth rate reduction and a correspondingly moderate compromise of pH homeostasis at pH 10.5 but increases the growth rate at pH 7.5 (Gilmour et al. manuscript submitted). A possible example of the “neutral pH as stress” scenario is autolysin activation in B. halodurans C-125 at the low end of its pH range for growth (Aono and Sanada, 1994).

Industrial Applications Enzymes Horikoshi (1991), (1996) has surveyed the applications of alkaliphiles in industrial processes, and Ito et al., 1998 have reviewed the alkaliphile enzymes that have been specifically used in laundry and dishwashing detergents. A major application of alkaliphile hydrolases, especially those that are also thermotolerant, is in laundry mixes that have alkaline pH. The recognition that these enzymes are useful fostered the rapid growth of information about alkaliphile proteases, in particular, but also of numerous additional hydrolases, including lipases, cellulases and pullulanases. Each of these classes of enzymes have other uses, such as the proteases in dehairing processes (Horikoshi, 1996) and in the degradation of gelatin-containing X-ray films for silver recapture (Fujiwara et al., 1991). Pullulanases and amylases also have had applications in other settings (e.g., the food industry).

CHAPTER 1.10

Alkaliphilic amylases and pullulanases have been isolated from numerous alkaliphiles, even including a haloalkaliphilic amylase from Natronococcus (Kobayashi et al., 1994). Alkaliphilic Bacillus sp. strain KSM-1378 was found to produce an alkaline amylopullulanase that has two independent active sites for the individual reactions (Ara et al., 1995). In some of the food industry applications, cold-adapted rather than heat-adapted enzymes are more useful, and such enzymes have accordingly been sought (Kimura and Horikoshi, 1990). Environmental concerns have created pressure to minimize the chlorine-intense processes used to bleach alkali-treated wood pulp. This, in turn, has encouraged the search for thermostable, alkaline xylanases that could substitute for the chemical process. Numerous xylanases have been characterized from alkaliphiles some of which were produced as exoenzymes and had desirable properties vis à vis the bleaching process, such as no cellulase activity (Nakamura et al., 1993; Nakamura et al., 1994; Blanco et al., 1995). Recent xylanase-producing alkaliphilic or alkalitolerant prokaryotes have been isolated from geothermal enrichments (Dimitrov et al., 1997; Lopez et al., 1998; Sunna et al., 1997). Horikoshi, 1991 also has noted reports of alkaliphilic enzymes that were being developed for possible use in the lignin-degradation processes that are important in pulping. Pectinases have been isolated from alkaliphiles and evaluated for use in fruit and vegetable processing industries, including the degradation of pectin in wastewater from such industries; some efficacy of these enzymes has been reported for these uses as well as for the retting process involved in production of Japanese paper (see Horikoshi, 1991). Recently, a pectate lyase, designated Pel-7, was purified from alkaliphilic Bacillus sp. strain KSM-P7 and characterized as a thermostable, highly alkaline enzyme (Kobayashi et al., 1999). Cyclomaltodextrin glucanotransferases (CGTases) produce cyclodextrins, which are homogeneous cyclic oligosaccharides, from starch. Cyclodextrins are used in industrial preparations of foods, pharmaceuticals and other chemicals. The application of CGTases from alkaliphiles has been an important application of alkaliphile enzymes (Horikoshi, 1991).

Antibiotic Production or Screening, Biotransformations Many conventional antibiotics are unstable at very alkaline pH values, but it is nonetheless possible that the alkaliphiles themselves (e.g., alka-

CHAPTER 1.10

liphilic Bacillus or actinomycetes) will produce alkali-stable antimicrobials. Among the published reports of such work are compounds isolated from alkaliphilic soil isolates (Sato et al., 1980; Tsujibo et al., 1992). It might be of particular interest to examine whether soda lake organisms, whose environment is more consistently and extremely alkaline, produce antibiotics. Since biological productivity is high, the capacity to produce antibiotics may well confer competitive advantage, and those antibiotics might have novel features of interest. As noted by Hsieh et al., 1998 in connection with Staphylococcus aureus mutants that lack a major multidrug efflux pump, even wild type cells will exhibit a 15- to 60-fold increase in sensitivity to antimicrobials at an alkaline pH that favors accumulation of cations and weak bases. Thus alkaliphiles are particularly susceptible to inhibition by toxic cations and weak bases and might offer a way to detect small quantities of such antibiotic substances in impure test samples unless the organisms are equipped with correspondingly high activity, multidrug efflux protection. The completion of the Bacillus halodurans C-125 sequence may clarify whether the latter adaptation is likely. Paavilainen and colleagues (Paavilainen et al., 1994; Paavilainen et al., 1995) have studied the dynamics of growth-induced changes in the medium pH and organic acid production by different alkaliphilic Bacillus species and have begun to characterize properties associated with alkaliphile catabolic patterns. When complete, the sequence of the B. halodurans C-125 genome may well provide clues for biotransformative and/or bioremediation capacities that will be of interest. Some heavy metal resistance determinants have been identified, for example, in alkaliphilic B. pseudofirmus OF4 (Ivey et al., 1992). Kimura et al., 1994 used a newly isolated alkaliphilic Bacillus to achieve conversions of hydroxyls of cholic acid to keto groups, in various combinations and at high yield.

Conclusions Fossil evidence suggests that extremely alkaliphilic bacteria probably were part of an ancient group of prokaryotes that evolved in natural enrichments. An enormous diversity exists among alkaliphiles. Thus, there may be a corresponding diversity in the alkaline adaptations both in presently existing soda lakes alkaliphiles and in the widespread alkaliphiles that may now be found even in apparently nonselective settings. Much of the exploitation of a considerable industrial potential and most of the studies of fundamental physiological adaptations have

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focused on alkaliphilic Bacillus species. The current trend of increasing inclusion of haloalkaliphilic archaea and different anaerobic alkaliphiles in these efforts is salutory. Much has been learned about the major features of pH homeostasis, motility, membrane transport, cell structure, protein adaptation to high pH, and conundrums such as oxidative phosphorylation; but each finding has also provoked new and interesting questions for which the contemporary explosion of genomics, and novel ecological, molecular, structural and biophysical approaches will provide the basis for new understanding and applications for alkaliphilic prokaryotes. Acknowledgments. Work from the author’s laboratory was supported in part by research grants GM28454 from the National Institutes of Health and DE-FG0286ER13559 from the Department of Energy. I am grateful to many colleagues in my laboratory and from other laboratories for sharing ideas and questions. I especially wish to express my appreciation to Arthur Guffanti, David Hicks, and Yi Wei, of Mount Sinai School of Medicine, and Masahiro Ito, of Tokyo University.

Literature Cited Aono, R., K. Horikoshi, and S. Goto. 1984. Composition of the peptidoglycan of alkalophilic Bacillus sp. 157. J. Bacteriol. 688–689. Aono, R. 1985. Isolation and partial characterization of structural components of the walls of alkalophilic Bacillus strain C-125. J. Gen. Microbiol. 131:105–111. Aono, R. 1989. Characterization of cell wall components of the alkalophilic Bacillus strain C-125: identification of a polymer composed of polyglutamate and polyglucuronate. J. Gen. Microbiol. 135:265–271. Aono, R., and M. Ohtani. 1990. Loss of alkalophily in cellwall-component-defective mutants derived from alkalophilic Bacillus C-125. Biochem. J. 266:933–936. Aono, R., M. Ito, and K. Horikoshi. 1992. Instability of the protoplast membrane of facultative alkaliphilic Bacillus sp. C-125 at alkaline pH values below the pH optimum for growth. Biochem. J. 285:99–103. Aono, R., H. Ogino, and K. Horikoshi. 1992. pH-dependent flagella formation by facultative alkaliphilic Bacillus sp. C-125. Biosci. Biotechnol. Biochem. 56:48–53. Aono, R., M. Ito, and K. Horikoshi. 1993. Occurrence of teichuronopeptide in cell walls of group 2 alkaliphilic Bacillus spp. J. Gen. Microbiol. 139:2739–2744. Aono, R., and T. Sanada. 1994. Hyper-autolysis of the facultative alkaliphile Bacillus sp. C-125 cells grown at neutral pH: culture-pH dependent cross-linking of the peptide moieties of the peptidoglycan. Biosci. Biotechnol. Biochem. 58:2015–2019. Aono, R. 1995. Assignment of facultatively alkaliphilic Bacillus sp. C-125 to Bacillus lentus group 3. Int. J. Syst. Bacteriol. 45:582–585. Aono, R., M. Ito, K. N. Joblin, and K. Horikoshi. 1995. A high cell wall negative charge is necessary for the growth

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Prokaryotes (2006) 2:309–335 DOI: 10.1007/0-387-30742-7_11

CHAPTER 1.11 ms i hpor tnyS

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Syntrophism among Prokaryotes BERNHARD SCHINK AND ALFONS J. M. STAMS

Introduction: Concepts of Cooperation in Microbial Communities, Terminology The study of pure cultures in the laboratory has provided an amazingly diverse diorama of metabolic capacities among microorganisms, and has established the basis for our understanding of key transformation processes in nature. Pure culture studies are also prerequisites for research in microbial biochemistry and molecular biology. However, desire to understand how microorganisms act in natural systems requires the realization that microorganisms don’t usually occur as pure cultures out there, but that every single cell has to cooperate or compete with other microor macroorganisms. The pure culture is, with some exceptions such as certain microbes in direct cooperation with higher organisms, a laboratory artifact. Information gained from the study of pure cultures can be transferred only with great caution to an understanding of the behavior of microbes in natural communities. Rather, a detailed analysis of the abiotic and biotic life conditions at the microscale is needed for a correct assessment of the metabolic activities and requirements of a microbe in its natural habitat. In many cases, relationships of bacteria with other organisms may be relatively unimportant, as appears to be the case with most aerobes: they can usually degrade even fairly complex substrates to water and carbon dioxide without any significant cooperation with other organisms. Nutritional cooperation may exist, but may be restricted to the transfer of minor growth factors, such as vitamins, from one organism to the other. However, we have to realize that this assumption is based on experience gained from pure cultures that were typically enriched and isolated in simple media, and the selection aimed at organisms that were easy to handle, independent of possible interactions with others. Estimations assume that we know only a small fraction of the microorganisms present in nature, perhaps 0.1–1.0%.

Thus, we cannot exclude that other bacteria out there might depend to a large extent on cooperation with partner microbes, and perhaps this is just one of the reasons why we failed so far to isolate them. Anaerobic microorganisms, on the other hand, depend to a great extent on the cooperation of several metabolic types of bacteria in feeding chains. The complete conversion of complex organic matter, e.g., cellulose, to methane and carbon dioxide in a lake sediment is catalyzed by the concerted action of at least four different metabolic groups of bacteria, including primary fermenters, secondary fermenters, and at least two types of methanogenic archaeobacteria (Bryant, 1979; McInerney, 1988; Stams, 1994; Schink, 1991; Schink, 1997). The degree of mutual dependence among these different metabolic groups (“functional guilds”) can vary considerably; whereas the latter members in the feeding line always depend on the former ones for substrate supply, they may also influence significantly the former chain members by removal of metabolic products. In an extreme case, this can mean that the fermenting bacterium depends entirely on cooperation with a methanogen to fulfill its function in, e.g., methanogenic fatty acid oxidation. This type of cooperation is called “syntrophic.” Mutual metabolic dependencies also can emerge from the cooperation of phototrophs with sulfur- or sulfate-reducing bacteria. Sulfurreducing, acetate-oxidizing, chemotrophic bacteria such as Desulfuromonas acetoxidans and phototrophic green sulfide-oxidizing bacteria like Chlorobium sp. can cooperate closely in a phototrophic conversion of acetate plus CO2 to bacterial cell mass, using a sulfide/sulfur cycle as an electron shuttle system between both. The two partners cooperate very closely also in this system for which the term “syntrophy” was originally coined (Biebl and Pfennig, 1978). Syntrophy is a special case of symbiotic cooperation between two metabolically different types of bacteria which depend on each other for degradation of a certain substrate, typically

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through transfer of one or more metabolic intermediate(s) between the partners. The pool size of the shuttling intermediate has to be kept low to allow efficient cooperation. The term “syntrophy” should be restricted to those cooperations in which partners depend on each other to perform the metabolic activity observed and in which the mutual dependence cannot be overcome by simply adding a cosubstrate or any type of nutrient. A classical example is the “Methanobacillus omelianskii” culture (Barker, 1940), which was later shown to be a coculture of two partner organisms, the S strain and the strain M.o.H. (Bryant et al., 1967). Both strains cooperate in the conversion of ethanol to acetate and methane by interspecies hydrogen transfer, as follows: Strain S: 2CH 3CH2 OH + 2H 2 O Æ 2CH 3COO- + 2H + + 4H 2 DG 0 ¢ = +19 kJ per 2 mol of ethanol Strain M.o.H. 4H 2 + CO2 Æ CH 4 + 2H 2 O DG 0 ¢ = -131kJ per mol of methane Corulture: 2CH 3CH2 OH + CO2 Æ 2CH 3COO- + 3H + + CH 4 DG 0 ¢ = -112kJ per mol of methane

Thus, the fermenting bacterium cannot be grown with ethanol in the absence of the hydrogenscavenging partner organism because it carries out a reaction that is endergonic under standard conditions. The first reaction can occur and provide energy for the first strain only if the hydrogen partial pressure is kept low enough ( 4-nitro (7.18) > 2-nitro (7.22) > 3,5dichloro (8.19) > 3-nitro (8.39) > 4-cyano (8.49) > 4-bromo (9.34; Tull and Withers, 1994). In retaining enzymes, the nucleophilic residue can be identified directly by trapping the intermediate with an appropriate inhibitor. Such inhibitors include model saccharides containing a fluorine substituent in the 2- or 5-position and a good leaving group, such as fluoride or dinitrophenolate (Williams and Withers, 2000). The substituted substrate forms a relatively stable covalent substrate-enzyme complex, involving the nucleophile residues. The complex is then subjected to proteolytic cleavage and sequencing of the glycosylated peptide. Recently, the use of protocols involving combined liquid chromatography and mass spectrometry has facilitated the identification of the modified residues. The acid-base residue in a retaining enzyme can be identified by a combination of kineticsbased methodologies. Mutation of this residue (usually to alanine) should affect the rate of both chemical steps, i.e., glycosylation and deglycosylation, though the effect on each step should be different. The effect on the glycosylation step will depend strongly on the leaving group ability of the aglycon. Thus, rates of hydrolysis for substrates with a poor leaving group should be affected much more strongly than those with a

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good leaving group. The deglycosylation step, however, will be affected equally for all substrates carrying different leaving groups, because the same glycosyl enzyme intermediate is hydrolyzed during this step. Thus, detailed kinetic analysis (i.e., determination of kcat and Km) with substrates bearing different leaving groups can reveal whether the corresponding mutation is the acid-base residue. It should be noted that this approach requires synthetic substrates that are not necessarily recognized by all families of enzymes and are not necessarily commercially available. For example, the family-11 xylanases fail to hydrolyze p-nitrophenyl xylobioside, which is an excellent substrate for the family-10 xylanases. The assignment of the acid-base catalyst can also be examined by use of external nucleophilic anions, such as azide. In this approach, termed “azide rescue,” the small azide anion enters the vacant space created by alanine replacement of the acidic amino acid residue. The azide reacts with the anomeric carbon instead of a water molecule to form the corresponding b-glycosyl azide product. In the absence of an acid-base catalyst, which normally provides general base catalysis during the second step, the deglycosylation step is severely affected. Thus, the acceleration of the reaction by the mutant enzymes in the presence of these external anions (provided that the second step is rate limiting) is a good indication that a mutant residue is the acid-base catalyst. Finally, the assignment of the acid-base catalyst can be tested by comparing the pH-dependence profiles for the wild-type and mutant enzymes. The profile for the native enzyme would approximate a perfect bell shape curve, reflecting the ionization of the two active site carboxylic acids, whereas the no reduction of activity at high pH values would be observed for the mutant. This pH dependency approach is also applicable for identifying the nucleophile residues and the catalytic residues in inverting enzymes.

Prokaryotic Cellulase Systems The cellulolytic bacteria produce a variety of different cellulases and related enzymes, which together convert the plant cell wall polysaccharides to simple soluble sugars that can subsequently be assimilated. The complement of cellulases and hemicellulases that are synthesized by a given bacterium for this purpose is referred to as its “cellulase system.” Different bacteria exploit different strategies for the ultimate degradation of their substrates. The given strategy is reflected by the complement and type(s) of enzymes produced by a given bacterium. The bacterial cellulase system may be char-

CHAPTER 1.19

acterized by free enzymes, cell-bound enzymes, multifunctional enzymes, cellulosomes, or any combination of the latter. Cellulase enzyme systems are comprised of several different types of components, each type may exist in a multiplicity of forms. To add to the complexity, the same component may exist as free individual entities in the culture fluid, as individual entities bound to cellulose, or associated with the cell surface. Alternatively, an individual component may be organized as part of a multicomponent cellulosome complex attached to the cell surface, to the cellulose, to both, or as free complexes in the culture fluid. Furthermore, the situation existing during growth under one set of conditions (e.g., pH, temperature, distribution of carbon source, etc.) may not exist under another, or may change considerably during the course of cultivation. The bacterium reacts to these changes and its production of cellulases and/or cellulosomes may reflect the dynamics of the growth conditions.

Free Enzymes As mentioned earlier in this chapter, the free enzymes in their simplest form comprise a catalytic module alone with no accessory domains or modules. Such enzymes often specialize in degrading soluble oligosaccharide breakdown products. Alternatively, such single-modular enzymes may rely on an intrinsic association with insoluble polysaccharide substrate such as cellulose, perhaps related to the active site of the enzyme. A higher order level of organization and activity are free enzymes composed of a polypeptide chain that includes both a catalytic domain together with a CBM. This basic bi-modular arrangement can be further extended by the inclusion of additional types of modules or repeating units of the same module, all of which serve to modulate the activity of the catalytic domain on the substrate. The intact free enzyme, however, remains unattached to other enzymes and can work in an independent manner on a given substrate.

Cell-Bound Enzymes Some enzymes are connected directly to the cell wall. In Gram-positive bacteria, this is frequently accomplished via a specialized type of module, the SLH (S-layer homology) module, previously shown to be associated with the cell surface of Gram-positive bacteria (Lupas et al., 1994). This arrangement may have evolved to provide a more economic degradation of insoluble substrates and to reduce competition with other bac-

Cellulose-Decomposing Bacteria and Their Enzyme Systems

teria for the soluble products, subject to diffusion in the media. As opposed to free enzymes, diffusion of an attached enzyme would itself be prevented. Examples of enzymes, which are bound to the cell surface via an SLH module include, a family5 cellulase and family-13 amylase-pullulanase from Bacillus, a family-10 xylanase from Caldicellulosiruptor (Saul et al., 1990), a family5 endoglucanase from Clostridium josui, a family-16 lichenase and family-10 xylanase from Clostridium thermocellum (Jung et al., 1998), and a variety of enzymes (family-10 xylanases, a family-5 mannanase and a family-13 amylasepullulanase) from different species of Thermoanaerobacter (Matuschek et al., 1996). The modular architecture of these enzymes may be particularly complicated, containing several different modules in a single polypeptide chain, thus forming extremely large enzymes sometimes comprising over 2,000 amino acids (Fig. 10).

Multifunctional Enzymes Some cellulases exhibit a more complex architecture in that more than one catalytic domain and/ or CBD may be included in the same protein. Examples of such enzymes are the very similar cellulases from Anaerocellum thermophilum (Zverlov et al., 1998) and Caldocellum saccharolyticum (Te’o et al., 1995), both of which contain a family-9 and a family-48 catalytic domain. Other paired catalytic domains include those from family 44 and either family 5 or 9. Such an arrangement might indicate a close cooperation between two particular catalytic domains, which may lead to synergistic action on the cellulosic substrate, thus portending on a smaller scale the advent of cellulosomes.

Cell-bound amylase-pullulanase from

X25 X25

X21

13

fn3

Thermoanaerobacter thermosulfurogenes

fn3

Fig. 10. A very large, cell-surface enzyme from Thermoanaerobacter thermosulfurogenes. The 1861-residue enzyme contains an SLH module, which is believed to mediate the attachment of the enzyme to the cell surface in Gram-positive bacteria. The enzyme contains a multiplicity of modules, which apparently serve to regulate the hydrolytic action of its single family-13 catalytic module with the complex substrate. Several X domains of unknown function may either represent as yet undescribed catalytic functions, carbohydratebinding activities or structural entities.

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Like the cellulases, xylanases also tend to exhibit a modular structure, being composed of multiple domains joined by linker sequences. Family-10 and -11 xylanases may be linked in the same polypeptide chain either to each other, to catalytic domains from families 5, 16 and 43 or to carbohydrate esterases (Flint et al., 1993; Laurie et al., 1997). One particularly interesting combination of multifunctional catalytic modules that appear in the same polypeptide chain is a typical xylanase together with a feruloyl esterase. Such a combination would allow the rapid cleavage of hemicellulose from the lignin in natural systems, i.e., the plant cell wall (see Fig. 3). In this manner, the xylan chain would be severed by the xylanase component (Xyn in Fig. 3) and the lignin-xylan association would be disconnected simultaneously by the feruloyl acid esterase (Fae in Fig. 3). Indeed, some xylanases are extremely complex in their modular architecture (Fig. 11). In addition to multiple catalytic modules, these enzymes often contain several different types of CBMs. Why would such a xylanase contain several types of CBM? And why would a xylanase contain a cellulose-specific CBD? Unlike the case of various cellulases, for which the CBD is usually essential for degrading insoluble crystalline cellulose, the CBMs of a hemicellulase do not necessarily bind the hemicellulose component (xylan). In some cases, its CBM is in fact an authentic CBD that situates the hemicellulase on the insoluble plant cell wall material by utilizing the most abundant and most stable cell-wall component—cellulose. Indeed, the three family-3 CBDs (CBM3) shown in Fig. 11 apparently bind to crystalline cellulose. Why would this xylanase require three tandem copies of the same type of CBD is yet another mystery that should eventually be addressed experimentally. At any rate, once bound via the

X31

CHAPTER 1.19

20

X32

X

SLH

X25-X25–X25–GH13–FN3-X31-FN3–CBM20–X32–X–SLH

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CHAPTER 1.19

Multi–modular Xylanase from Caldicellulosiruptor

22

22

10

3

3

3

43

6

CBM22-CBM22-GH10-CBM3-CBM3-CBM3-GH43-CBM6

cellulose component of the plant cell wall composite substrate, the immobilized enzyme then acts on the accessible and appropriate hemicellulose components. Once thus situated on the plant cell wall, another type of CBM on the same molecule would then assist in the binding to the xylan (or mannan, etc.) component to direct the appropriate catalytic module to its true substrate. Hence, the modular proximity of the xylanase shown in Fig. 11 would presumably indicate that the two CBM22s would modulate the action of the family-10 catalytic module, and the C-terminal CBM6 would facilitate the catalysis by the family-43 module. Together, the two catalytic modules would act synergistically to degrade susceptible plant cell wall components. In this context, the complex architecture of a xylanase would reflect the complex chemistry of its substrate and the neighboring polymers of its immediate environment in the plant cell wall.

Cellulosomes Cellulosomes are multienzyme complexes, which bind to and catalyze the efficient degradation of cellulosic substrates. The first cellulosome was discovered while studying the anaerobic thermophilic bacterium, Clostridium thermocellum (Bayer et al., 1983; Lamed et al., 1983). Since its initial description in the literature, the cellulosome concept has been subject to numerous reviews (Bayer et al., 1996; Béguin and Lemaire, 1996; Belaich et al., 1997; Doi et al., 1994; Doi and Tamura 2001; Felix and Ljungdahl, 1993; Karita et al., 1997; Lamed and Bayer, 1988; Lamed and Bayer, 1991; Lamed and Bayer, 1993; Lamed et al., 1983; Shoham et al., 1999). Cellulosomes in C. thermocellum exist in both cell-associated and extracellular forms, the cell-

Fig. 11. A very large, multimodular xylanase from Caldicellulosiruptor. The 1,795-residue enzyme contains 8 separate modules, including 2 catalytic modules from families 10 (invariably a xylanase) and 43 (frequently an arabinofuranosidase). These are modulated by numerous carbohydratebinding modules, which include 3 from family 3 (likely for binding to crystalline cellulose), 2 from family 22 (newly classified and shown to function in xylan binding and one from family 6.

associated form being associated with polycellulosomal protuberance-like organelles on the cell surface. Later, cellulosomes were detected in other cellulolytic organisms (Lamed et al., 1987; Mayer et al., 1987), including Acetivibrio cellulolyticus, Bacteroides cellulosolvens, Clostridium cellulovorans and Ruminococcus albus, all of which contained protuberance-like organelles on their surfaces (Bayer et al., 1994; Lamed and Bayer, 1988; Fig. 12). The cellulosomes contain numerous components, many of which were shown to display enzymatic activity. They also contain a characteristic nonenzymatic high-molecular-weight component. This component proved to be highly antigenic and glycosylated (Bayer et al., 1985). The cellulosomal enzymatic subunits from this organism showed a broad range of different cellulolytic and xylanolytic activities (Morag et al., 1990). Ultrastructural evidence indicated the multisubunit nature of the cellulosome (Fig. 13). Eventually, genetic engineering techniques led to the sequencing of cellulosomal genes in C. thermocellum and several other bacteria, thus confirming the existence of cellulosomes as a major paradigm of prokaryotic degradation of cellulose and related plant cell wall polysaccharides.

Clostridium Thermocellum Cellulosomal Subunits and Their Modules A simplified schematic view of the cellulosome from C. thermocellum and its interaction with its substrate is shown in Fig. 14. The cellulosomal enzyme subunits were found to be united into a complex by means of a unique class of nonenzymatic, multimodular polypeptide subunit, termed “scaffoldin” (Bayer et al., 1994). The

CHAPTER 1.19

Cellulose-Decomposing Bacteria and Their Enzyme Systems A

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B

Fig. 12. Scanning electron microscopy (SEM) of Acetivibrio cellulolyticus showing the presence of large characteristic protuberance-like structures on the cell surface. Cells are shown in the free state (A) or bound to cellulose (B). Cell preparations were treated with cationized ferritin before processing. Cationized ferritin has been shown to stabilize such surface structures, thus allowing their ultrastructural visualization (Lamed et al., 1987a; Lamed et al., 1987b). Without pretreatment with cationized ferritin, these structures are invisible. In (B), the cellulose-bound cells appear to be connected to the substrate via structural extensions of the cell-surface protuberances. Such a mechanism was originally observed for other cellulolytic prokaryotes, e.g., C. thermocellum (Bayer and Lamed, 1986).

50 nm Fig. 13. Comparison between negative staining (bottom) and cryo images (top) of the purified cellulosome from C. thermocellum, adsorbed on cellulose microcrystals from the algae, Valonia ventricosa. The images illustrate the diversity of shapes of the cellulosomes, which adopt either compact or loosely organized ultrastructure. In the cryo images, the subunits of the cellulosomes (i.e., the individual enzymatic components) are clearly visible. Micrographs courtesy of Claire Boisset and Henri Chanzy (CNRS—CERMAV, Grenoble, France).

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CHAPTER 1.19

The cellulosome of C.thermocellum

Cellulosome

Cell

Enzymatic subunits Key

CBD

Anchoring protein

Type-I cohesin domain Type-II dockerin domains

Scaffoldin subunit

Catalytic domain Type-I dockerin domains Type-II cohesin domain SLH module

Cellulose

Cellulose chain

Fig. 14. Simplified schematic view of the molecular disposition of the cellulosome and one of the associated anchoring proteins on the cell surface of C. thermocellum. The key defines the symbols used for the modules, from which the different cellulosomal proteins are fabricated. The progression of cell to anchoring protein to cellulosome to cellulose substrate is illustrated. The SLH module links the parent anchoring protein to the cell. The cellulosomal scaffoldin subunit performs three separate functions, each mediated by its resident functional domains: 1) its multiple type-I cohesins integrate the cellulosomal enzymes into the complex via their resident type-I dockerins, 2) its family-IIIa CBD binds to the cellulose surface, and 3) its type-II dockerin interacts with the type-II cohesin of the exocellular anchoring protein.

scaffoldins usually contain a family-3 CBD that provides the cellulose-binding function. The scaffoldins also contain multiple copies of a definitive type of module, called “the cohesin domain.” The cellulosomal enzyme subunits, on the other hand, contain a complementary type of module, called “the dockerin domain.” The interaction between the cohesin and dockerin domains provides the definitive molecular mechanism that integrates the enzyme subunits into the cellulosome complex (Salamitou et al., 1994; Tokatlidis et al., 1991; Tokatlidis et al., 1993). Cohesin and dockerins are considered to be cellulosome “signature sequences”—i.e., their presence is a good indication of a cellulosome in a given bacterium (Bayer et al., 1998). The major difference between free enzymes and cellulosomal enzymes is that the free enzymes usually contain a CBD for guiding the catalytic domain to the substrate, whereas the cellulosomal enzymes carry a dockerin domain that incorporates the enzyme into the cellulosome complex. Otherwise, both the free and cellulosomal enzymes contain very similar types

of catalytic domains. The cellulosomal enzymes rely on the Family-3a CBD of the scaffoldin subunit for collective binding to crystalline cellulose. The incorporation of the multiplicity of enzyme subunits into the cellulosome complex is a function of the repeated copies of the cohesin module borne by the scaffoldin subunit. For most species of scaffoldin, the cohesins have been classified as type-I on the basis of sequence homology. The cohesin module is composed of about 150 amino acid residues. The basic structure of the cohesin is known and comprises a ninestranded b sandwich with a jelly-roll topology (Shimon et al., 1997; Spinelli et al., 2000; Tavares et al., 1997). The dockerin domain contains about 70 amino acids and is distinguished by a 22-residue duplicated sequence (Chauvaux et al., 1990), which bears similarity to the well-characterized EFhand motif of various calcium-binding proteins (e.g., calmodulin and troponin C). Within this repeated sequence is a 12-residue calciumbinding loop, indicating that calcium-binding is an important characteristic of the dockerin

CHAPTER 1.19

Cellulose-Decomposing Bacteria and Their Enzyme Systems

domain. This assumption was eventually confirmed experimentally (Yaron et al., 1995). The specificity characteristics of the cohesin-dockerin interaction also have been investigated. The results showed that four suspected residues may serve as recognition codes for interaction with the cohesin domain (Mechaly et al., 2000; Mechaly et al., 2001; Pagès et al., 1997). The three-dimensional solution structure of the 69residue dockerin domain of a Clostridium thermocellum cellulosomal cellulase subunit was recently determined (Lytle et al., 2001). As predicted earlier (Bayer et al., 1998; Lytle et al., 2000; Pagès et al., 1997), the structure consists of two Ca2+-binding loop-helix motifs connected by a linker; the E helices entering each loop of the classical EF-hand motif are absent from the dockerin domain. The scaffoldin of C. thermocellum also contains a special type of dockerin domain. This dockerin failed to bind to the cohesins from the same scaffoldin subunit, but instead interacted with a different type of cohesin—termed “typeII cohesins”—identified on the basis of sequence homology (Salamitou et al., 1994). These cohesins are somewhat different than those of type I, having an additional segment and diversity in the latter half of the sequence. The type-II cohesins were discovered as component parts of a group of noncatalytic cell-surface “anchoring” proteins on C. thermocellum (Leibovitz and Béguin, 1996; Leibovitz et al., 1997; Lemaire et al., 1995; Salamitou et al., 1994). The three known anchoring proteins in C. thermocellum contain different copy-numbers of the type-II cohesins as illustrated in Fig. 15. Each of these anchoring proteins also contains an S-layer homology (SLH) module, analogous to those of the cell-bound enzymes mentioned above. The intervening sequences, however, between the cohesins and SLH domains are different. In any case, the typeII cohesins selectively bind the type-II dockerins, and the cellulosome (i.e., the scaffoldin subunit together with all of its enzyme subunits) is thereby incorporated into the cell surface of C. thermocellum.

Similarity and Diversity of Scaffoldins from Different Species The modular architecture of the known scaffoldins and their comparison to that of Clostridium thermocellum is presented in Fig. 16. Two new scaffoldins have recently been described for Acetivibrio cellulolyticus and Bacteroides cellulosolvens that, like C. thermocellum, carry dockerin domains at their C terminus (Ding et al., 1999; Ding et al., 2000). The A. cellulolyticus genome also includes a gene (immediately downstream of the scaffoldin gene) that contains type-II

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Outer-layer proteins of C. thermocellum involved in anchoring the cellulosome onto the cell surface

SdbA

Orl2p

OlpB

Key Type II cohesin domain SLH module

Fig. 15. Schematic representation of the known anchoring proteins of the C. thermocellum cell surface. Each protein bears an SLH domain that connects the protein to the cell surface via yet undefined surface components. The different proteins carry different numbers of type-II cohesins. SdbA has one cohesin, Orf2p has 2 and OlpB has 4, presumably allowing the corresponding number of scaffoldins (i.e., cellulosomes) to be attached to the given protein.

cohesins that may represent an anchoring protein. It thus seems that the arrangement of the cellulosome on the cell surface of these latter strains may be analogous to that of C. thermocellum. It is interesting to note that the cohesins of the Bacteroides cellulosolvens scaffoldin are clearly type-II cohesins and not of type I. This infers that there is not a clear linkage between the type-II cohesins and anchoring proteins. The scaffoldins from the other clostridial species thus far described all lack “type-II dockerin” domains, the inference being that cells of C. cellulovorans, for example, would apparently not bear anchoring proteins that contain type-II cohesins. It thus follows that either their cellulosomes are not surface bound or, if indeed they are surface components, then their anchoring thereto is accomplished via an alternative molecular mechanism. Recently (Doi and Tamura, 2001; Tamaru and Doi, 1999a; Tamaru et al., 1999b), a cell-surface binding function has been proposed for a domain of unknown function, designated “X2” (Coutinho and Henrissat, 1999b; Coutinho and Henrissat, 1999c) of the scaffoldin from C. cellulovorans. On the basis of sequence alignment of a few conserved identical amino acids with S-layer proteins from Mycoplasma hyorhinis and Plasmodium reichenowi, the authors consider that this domain may be recognized as an SLH domain. The four X2 domains of the C. cellulovorans scaffoldin are very similar in sequence to the X-domains from the scaffoldins of C. cellulolyticum and C. josui,

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CHAPTER 1.19

Classification of scaffoldins from different species

Class I Scaffoldins

Clastridum thermacellum

Class I Scaffoldins

Ruminococcal Scaffoldins

Clastridum cellulovorans Ruminocacus flavefaciers

Clastridum cellulolytum Acetiwbrio cellulolyticus

Clastridum josul Bocteroides cellulosalvens

Fig. 16. Schematic view of the modular similarity and diversity of scaffoldins from different cellulosome species. Class-I scaffoldins feature an internal CBD and a C-terminal type-II dockerin domain. Class-II scaffoldins exhibit an N-terminal CBD and lack a dockerin domain. The newly described scaffoldins from Ruminococcus flavefaciens lack a defined CBD. The functional role of the two different X domains in the two R. flavefaciens scaffoldins is currently unknown. All of these scaffoldins contain multiple copies of cohesin domains.

which contain only two and one copies of this domain, respectively. If this domain functions in attaching the scaffoldin with its complement of enzymes to the cell surface, it is unclear why there would be different copy numbers of the domain in the different scaffoldins. Likewise, one of the C. cellulovorans cellulosomal enzyme components (EngE) also contains a triplicated segment of unknown function, designated “X48” (Coutinho and Henrissat, 1999b; Coutinho and Henrissat, 1999c) that the authors consider to be involved in cell-surface attachment (Tamaru and Doi, 1999a). In any case, final proof of the function of the X2 and X48 domains awaits biochemical examination, as has been clearly achieved for the SLH domain of the C. thermocellum anchoring proteins (Chauvaux et al., 1999; Lemaire et al., 1998). Finally, two new scaffoldins have recently been sequenced from the rumen bacterium, Ruminococcus flavefaciens (Ding et al., 2001). Although each of the two proteins contains multiple cohesins, their sequences indicate that they are neither

of type-I or type-II, but occupy their own phylogenetic branch. Interestingly, the ruminococcal scaffoldins lack a known type of CBD. Both have dissimilar X domains of unknown function, the sequences of which bear no resemblance to any other known module. Both X domains were expressed, but the resultant proteins failed to bind to cellulose. The lack of a scaffoldin CBD raises the question as to how the ruminococcal cellulosome(s) and/or the bacterium bind to the substrate. Perhaps it does so like another closely related species, R. albus, which binds cellulose via a noncellulosomal cell-surface protein (Pegden et al., 1998).

Schematic Comparison of Prokaryotic Cellulase Systems In this section, we will describe schematically the similarity and diversity of representative enzyme systems, demonstrating different strategies, from

CHAPTER 1.19

Cellulose-Decomposing Bacteria and Their Enzyme Systems

different plant cell wall degrading bacteria. It is emphasized that the accumulating information is based on what is known currently from biochemical data combined with gene sequencing and bioinformatics. The information is still rather sketchy but quite revealing when compared among different bacteria. As time progresses and the entire genomes of cellulolytic microorganisms become known, the data concerning the complement of enzymes produced by a given bacterium will be complete, and we will be able to speculate with heightened certainty how the various cellulase systems might have evolved. A survey of genes, however, does not inform us how a given bacterial system is regulated and what role(s) the bacterium and its enzyme system may play in nature. The explosive development of molecular biology techniques, however revealing, cannot supplant the fundamental contribution of biochemical and ecological approaches to the study of microbial degradation of cellulose and other plant cell wall polysaccharides.

Free Enzyme Systems Many cellulolytic microorganisms show a very similar pattern in the types of enzymes that comprise the complement of their cellulase system. For the purposes of this discussion, the concept of “cellulase system” will include the complement of all plant cell wall hydolyzing enzymes and other glycosyl hydrolases, including the different cellulases per se, the hemicellulases (e.g., xylanases and mannanases), etc. The cellulase system of the mesophilic cellulolytic aerobe, Cellulomonas fimi, is one of the first studied, and has since been one of the most studied bacterial cellulase systems (O’Neill et al., 1986; Shen et al., 1995; Whittle et al., 1982). The enzymes of this bacterium are essentially free enzymes, which allowed their early isolation and characterization. Moreover, the genes of the cellulases from this bacterium were of the earliest to have been sequenced. To date, about 10 glycosyl hydrolases have been sequenced from Cellulomonas fimi. Their modular composition and family associations are shown symbolically in Fig. 17. As an example of a free enzyme system, most of the enzymes bear a substrate-targeting CBM—in this bacterium, most of the CBMs are from family 2. Several of the enzymes have multiple copies of the fibronectin 3 (FN3) domain, the function of which is still unknown. The Cellulomonas system includes two family6 enzymes—an endoglucanase and an exoglucanase (cellobiohydrolase) of the types described in Fig. 4. The modularity of the endoglucanase is

603

very simple, having the family-6 catalytic module together with a family-2 CBM. The cellobiohydrolase is a bit more complex with three additional FN3 domains that separate the same two types of modules. Another cellobiohydrolase (that exhibits processive cleavage of the substrate) is from family 48. Its general modular architecture is similar to that of the family-6 cellobiohydrolase with the substitution of the catalytic module from a different family. The cellulase system from this organism also includes two family-9 cellulases with modular themes B and D, familiar to us from the earlier description (Fig. 7). In addition, a simple family-5 cellulase and an interesting cell-borne family-26 mannanase are components of the system. The fact that an enzyme bears an SLH domain and is presumably cell-associated would underscore its importance to the cell. Finally, three xylanases are currently known for Cellulomonas fimi. One of these xylanases is a simple enzyme consisting of a family-10 catalytic domain connected to a family-2 CBM. The other two are more complicated, each containing two catalytic domains— either a family-10 or -11 domain and a carbohydrate esterase (in both cases, probably an acetyl xylan esterase; Fig. 3)—plus several CBMs. This rather complex system is probably not nearly complete, and more enzymes will inevitably be described in the future. A second example of a free enzyme system, from the aerobic thermophilic bacterium Thermobifida fusca (formerly classified as Thermomonospora fusca), has also been studied extensively (Wilson, 1992; Wilson and Irwin, 1999). A brief comparison of its known enzyme components (Fig. 18) shows a striking resemblance to those of Cellulomonas (compare Figs. 17 and 18). According to known data, both species produce similar types of cellulases from families 5, 6, 9 and 48 plus xylanases from families 10 and 11. Nevertheless, the modular repertoire of the corresponding enzyme in T. fusca is generally somewhat simpler. For example, two of the T. fusca cellulases include single FN3 domains, whereas several Cellulomonas cellulases harbor multiple copies of the same domain. Some T. fusca enzymes lack accessory modules other than a cellulose-binding CBM, whereas the corresponding Cellulomonas enzyme is elaborated by multiple copies of accessory modules. In some cases though, the respective CBMs appear on opposite termini of the polypeptide chain (i.e., the family-48 and family-5 cellulases). The complement of enzymes and their modular content of the free enzyme systems from Cellulomonas and T. fusca are not necessarily similar in other free enzyme systems. Many free enzyme systems, such as those of Butyrivibrio fibrisolvens, Pseudomonas fluorescens, Fibro-

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E.A. Bayer, Y. Shoham and R. Lamed

CHAPTER 1.19

6

Cel6B

6

CBM2-GH6

fn3

2

fn3

Cel6A

fn3

Known Enzymes from Cellulomonas fimi

2

GH6-FN3-FN3-FN3-CBM2 48

GH9-CBM3c-FN3-FN3-FN3-CBM2

4

X

I9

Cel9B

9 4

X

GH48-FN3-FN3-FN3-CBM2

Xyn10A (Cex)

fn3

fn3

5

GH10-CBM2

Xyn10B (Xync)

2

GH5-FN3-FN3-CBM2

ManA

23

26

Ce4

22

9

10

X

9

CE4-CBM22-GH10-CBM9-CBM9-X

Xyn11A

X

SLH

2

10

CBM4-CBM4-Ig-GH9-X-X

Cel5A

fn3

Cel48A

fn3

2

fn3

fn3

fn3

3c

Cel9A

fn3

9

11

2

2

Ce4

GH11-CBM2-CE4-CBM2

GH26-CBM23-SLH-X

Fig. 17. Cellulomonas fimi cellulase system: Symbolic view of the enzyme components and their modular architecture. An example of a cell-free enzyme system. The modular content of the enzymes in this and subsequent figures is shown from (left to right) the N-terminus to the C-terminus of the polypeptide chain. The family numbers of the given domains are enumerated, the catalytic modules given in red. Key to symbols: GH, glycosyl hydrolase (e.g., cellulase, xylanase and mannanase); CE, carbohydrate esterase (e.g., acetyl xylan esterase and ferulic acid esterase); CBM, carbohydrate-binding module (e.g., CBD, cellulose-binding domain); SLH, S-layer homology (domain); FN3, fibronectin-3 (domain); Ig, immunoglobulin-like domain; and X, domain of unknown function.

Known Enzymes from Thermobifida fusca

6

2

Cel6B(E3)

6

Cel6A(E2)

Cel9A(E4)

fn3

CBM2-GH6

3c

GH6-CBM2

Cel48A(E6)

2

2

Ig

fn3

9 4

2

Xyn10A

2

5

X

10

2

GH10-CBM2

CBM4-Ig-GH9-FN3-CBM2

Cel5A(E5)

X

48

CBM2-X-X-GH48

GH9-CBM3c-FN3-CBM2

Cel9B(E1)

2

Xyn11A

CBM2-GH5

bacter succinogenes, various species of Streptomyces, Erwinia and Thermatoga, appear to have several cellulases, xylanases and mannanases from the common families, together with other glycosyl hydrolases, e.g., arabinosidases, lichenases, amylases, pullulanases, galactanases, polygalacturonase, glucuronidases and pectate lyases. In many of these bacterial enzymes, the family-2 CBM appears to predominate as a common

11

2

GH11-CBM2

Fig. 18. Thermobifida fusca cellulase system. A cell-free enzyme system. The modular content of the enzymes is shown from (left to right) the Nterminus to the C-terminus of the polypeptide chain. Compare with the Cellulomonas system (Fig. 17). Key to symbols: GH, glycosyl hydrolase (e.g., cellulase, xylanase and mannanase); CBM, carbohydrate-binding module (e.g., CBD, cellulose-binding domain); FN3, fibronectin-3 (domain); Ig, immunoglobulin-like domain; and X, domain of unknown function.

cellulose-binding domain, but in others (e.g., Erwinia) relevant enzymes usually bear a cellulose-binding CBM from family-3. Nevertheless, in many of the free systems, many enzymes are characterized by CBMs from other families as well as other noncatalytic domains of unknown function (X domains). Once again, until the genome sequences of cellulolytic prokaryotes are widely available, we are still lim-

CHAPTER 1.19

Cellulose-Decomposing Bacteria and Their Enzyme Systems

ited in our capacity to compare among the enzyme systems because our knowledge of their enzyme sequences is incomplete.

605

enzymes is usually one or more copies of a family-3 CBM. Other bacterial strains that include at least one free bifunctional enzyme in their enzyme systems are Anaerocellum thermophilum, Bacillus stearothermophilus, Fibrobacter succinogenes, Prevotella ruminicola, Ruminococcus albus, Ruminococcus flavefaciens, Streptomyces chattanoogensis and the thermophilic anaerobe NA10. Unlike the Caldicellulosiruptor system, most of the free bifunctional enzymes in the latter strains appear to be isolated cases in the given system, rather than being a common character of their enzymes.

Multifunctional Enzyme Systems In an extremely thermophilic bacteria, classified as Caldicellulosiruptor, the enzymes currently characterized in this system also appear to be free enzymes, but their modular organization is of a higher order (Daniel et al., 1996; Gibbs et al., 2000; Reeves et al., 2000). Many of the enzymes of this system are bifunctional in that they contain two separate catalytic modules in the same polypeptide chain (Fig. 19). As mentioned earlier, the appearance of two catalytic modules in the same enzyme would infer a distinctive synergistic action between the two. Thus, in CelA, the family-9 and -48 catalytic modules would be expected to work in concerted fashion on crystalline cellulose. In another type of enzyme, the family-10 xylanase and family-5 cellulase would likely be most effective on regions of the plant cell wall that are characterized by cellulose-xylan junctions. The diversity in the modular architecture of the family-10 xylanases is particularly striking, and the various combinations of this type of catalytic module are apparently important to the sustenance of the bacterium in its environment. One of these xylanases appears to be attached to the cell surface via SLH domains. In contrast to the Cellulomonas and T. fusca enzymes that often harbor a family-2 CBM, the module responsible for binding to cellulosic substrates in Caldicellulosiruptor

Cellulosomal Systems The inclusion of enzymes into a cellulosome via the noncatalytic scaffoldin subunit represents a higher level of organization. The association of complementary enzymes into a complex is considered to contribute sterically to their synergistic action on cellulose and other plant cell wall polysaccharides. As mentioned earlier, in the case of Clostridium thermocellum, Acetivibrio cellulolyticus and Bacteroides cellulosolvens, the cellulosomes appear to be attached to the cell surface. The cellulosomes of C. cellulolyticum, C. cellulovorans and C. josui may also be cellassociated, but if so, the lack of a scaffoldinborne dockerin and reciprocal anchoring protein would suggest an alternative mechanism. The cellulosomes of C. cellulolyticum, C. cellulovorans and C. josui are very similar. The genes encoding for many or most of the enzymes in all

Known Enzymes from Caldicellulosiruptor

CelA

9

3c 3

48

3

10

XynA

GH10

GH9-CBM3c-CBM3-CBM3-GH48

Man-EG’ase

Fig. 19. Caldicellulosiruptor enzyme system. An example of a cell-free enzyme system that includes several multifunctional enzymes. The modular content of the enzymes is shown from (left to right) the N-terminus to the C-terminus of the polypeptide chain. Key to symbols: GH, glycosyl hydrolase (e.g., cellulase, xylanase and mannanase); CBM, carbohydrate-binding module (e.g., CBD, cellulose-binding domain); and SLH, S-layer homology (domain).

5

3

44

3

XynE & XynI

GH5-CBM3-CBM3-GH44

Cel

5

3

10

XynB

10

3

3

3

5

GH10-CBM3-CBM3-CBM3-GH5

XynF (Abn) 43

X

43

GH43-X-GH43

22

10

22

22

10

9

9

X

SLH

CBM22-CBM22-GH10-CBM9-CBM9-X-SLH

GH10-CBM3-GH5

CelB

22

CBM22-CBM22-GH10

XynC

22

22

10

3

3

3

43

6

CBM22-CBM22-GH10-CBM3-CBM3-CBM3-GH43-CBM6

XynD

11

9

GH11-CBM9

E.A. Bayer, Y. Shoham and R. Lamed

three cellulosomal systems are arranged in a large cluster on the chromosome. Some of the cellulosomal genes, however, are located outside of the cluster in other regions of the chromosome. The majority of the cellulosome gene clusters from C. cellulolyticum and C. cellulovorans have been sequenced (Bagnara-Tardif et al., 1992; Belaich et al., 1999; Tamaru et al., 2000b). In contrast, the cellulosomal genes from C. thermocellum are generally scattered over a large portion of the chromosome (Guglielmi and Béguin, 1998). A few small clusters of cellulosomal genes are apparent in the genome, including a scaffoldin-containing cluster that also contains several cell-surface anchoring proteins (Fujino et al., 1993). The following descriptive analysis serves to compare the cellulosomal system of these three microorganisms. Cellulosomal components from Clostridium cellulolyticum. All of the sequenced enzymes from this organism are relatively common cellulases (Belaich et al., 1999). None of the known cellulosomal enzymes yet described for this species contains more than one catalytic module (Fig. 20). The largest one, CelE (estimated at 94 kDa), is a theme-D family-9 cellulase (Gaudin et al., 2000). The critical family-48 cellulase (CelF) is also a major cellulosome component (Reverbel-Leroy et al., 1997). Interestingly, the gene cluster of C. cellulolyticum contains three copies of other family-9 cellulases (CelG, CelH and CelJ), all of which contain the themeB fused family-3c CBM (Belaich et al., 1998; Fig. 8). The currently known cellulosome system in this bacterium also contains two family-5 cellulases (CelA and CelD), a family-5 mannanase (ManK, which bears an N-terminal rather than C-terminal dockerin) and a family-8 cellulase (CelC). Biochemical characterization of the C. cellulolyticum cellulosome demonstrated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) a 160-kDa scaffoldin band and up to 16 smaller bands, representing putative enzyme subunits (Gal et al., 1997). Many of these were clearly identified as known gene products. Only two cellulosomal cellulase genes are currently known to be located outside of the gene cluster. Further work on the enzyme system of this species may yet provide more complicated multimodular enzymes and/or other types of enzymes, such as hemicellulases. In this context, recent biochemical evidence has suggested that xylanases from C. cellulolyticum are also organized in a cellulosome-like complex, but defined xylanase sequences are still lacking from this organism (Mohand-Oussaid et al., 1999). The known activity of this organism on other plant cell wall

CHAPTER 1.19 Known Enzymes from Clostridium cellulolyticum Cellulosomal Enzymes 9

CelE

Ig

606

4

CBM4-Ig-GH9-Doc

CelA

GH5-Doc

48

CelF

GH48-Doc

CelD

9

GH9-CBM3c-Doc

5

GH5-Doc

3c

CelG

5

Mank

5

9

CelH 9

CelJ

Doc-GH5

3c

GH9-CBM3c-Doc

CelC

3c

8

GH8-Doc

GH9-CBM3c-Doc

Scaffoldin

3a

CBM3a-X-7Coh-X-Coh

Fig. 20. Clostridium cellulolyticum enzyme system. An example of a cellulosomal system. The modular content of the enzymes is shown from (left to right) the N-terminus to the C-terminus of the polypeptide chain. Key to symbols: GH, glycosyl hydrolase (e.g., cellulase, xylanase and mannanase); CBM, carbohydrate-binding module (e.g., CBD, cellulosebinding domain); and Doc, dockerin domain.

polysaccharides would indicate that numerous other enzymes, either cellulosomal or not, remain as yet undiscovered. Cellulosomal components from Clostridium cellulovorans. Like C. cellulolyticum, the cellulases from this organism are relatively simple (Fig. 21). In addition to the cellulosomal enzymes thus described, at least three noncellulosomal endoglucanases have also been partially or totally sequenced (Doi et al., 1998; Tamaru et al., 1999b). Several of the cellulosomal enzymes are architecturally synonymous to those of the C. cellulolyticum system (compare Figs. 20 and 21). This includes the critical family-48 cellulase (ExgS; Liu and Doi, 1998), two copies of the theme-B family-9 cellulase (EngH and EngY), a family-5 endoglucanase and a family-5 mannanase that bears an N-terminal dockerin (Tamaru and Doi, 2000a). Rather than a single theme-D family-9 cellulase as in C. cellulolyticum, the C. cellulovorans system contains two such enzymes (EngK and EngM). The C. cellulovorans cellulosome also appears to contain an unusual theme-A

CHAPTER 1.19

Cellulose-Decomposing Bacteria and Their Enzyme Systems

607

Known Enzymes from Clostridium cellulovorans Cellulosomal Enzymes Non-Cellulosomal 5 EngE

EngY 3x48–GH5–X–Doc

EngA GH9–(?)

9 EngL

4

GH9–Doc

GH5–Doc

CBM4–Ig–GH9–Doc 48

5

2

5

EngB

4

EngD

GH5–CBM2

9

EXgS

Enzymes 9

CBM4–Ig–GH9–Doc

EngM

3c

X17–GH9–CBM3c–Doc

9 EngK

9

x7

5

17

GH5–CBM17

5

ManA

Cel5A (EngF)

Doc–GH5

GH48–Doc 9 3c

EngH

LyaA

GH9–CBM3c–Doc

Lya4–Doc

Scaffoldin x

x x

x 3a CBM3a–X–2Coh–X–6Coh–2X–Coh Fig. 21. Clostridium cellulovorans: A second cellulosomal system. The modular content of the enzymes is shown from (left to right) the N-terminus to the C-terminus of the polypeptide chain. Key to symbols: GH, glycosyl hydrolase (e.g., cellulase, xylanase and mannanase); CBM, carbohydrate-binding module (e.g., CBD, cellulose-binding domain); Doc, dockerin domain; SLH, S-layer homology (domain); Ig, immunoglobulin-like domain; and X, domain of unknown function.

family-9 cellulase (EngL) that lacks helper domains. The remaining two known cellulosomal enzymes are thus far unique to C. cellulovorans. A dockerin-bearing pectate lyase (LyaA) infers that the bacterium would degrade pectin (Tamaru and Doi, 2001). Indeed, early evidence (Sleat et al., 1984) indicated that, in addition to cellulose, C. cellulovorans is capable of assimilating a wide variety of other plant cell wall polysaccharides, including, xylans, pectins and mannans. As in the case of C. celluolyticum, it seems that future work will yield new sequences of many other types of cellulosomal and noncellulosomal enzymes.

More significant to the cellulosomal system of C. cellulovorans, perhaps, is the large family-5 enzyme that purportedly comprises both an Nterminal SLH domain and a C-terminal dockerin (Tamaru and Doi, 1999a). This arrangement may imply that the entire cellulosome is bound to the cell surface via this enzyme. If this proves to be the case, it is interesting to speculate whether the C. cellulolyticum and C. josui cellulosomes are also connected to the cell surface by a similar, but as yet undiscovered enzyme that bears both SLH and dockerin domains. Cellulosomal components from Clostridium thermocellum. Compared to the cellulosomal

608

E.A. Bayer, Y. Shoham and R. Lamed

systems of C. cellulovorans and C. cellulolyticum, the enzymes from C. thermocellum are relatively large proteins, ranging in molecular size from about 40–180 kDa (Bayer et al., 1998; Bayer et al., 2000; Béguin and Lemaire, 1996; Felix and Ljungdahl, 1993; Lamed and Bayer, 1988; Shoham et al., 1999). Examination of Fig. 22 reveals why these enzymes are so big—many of the larger ones contain multiple types of catalytic domains as well as other functional modules as an integral part of a single polypeptide chain (see Table I in Bayer et al., 1998, for a list of relevant references). In addition to the cellulosomal enzymes, several noncellulosomal enzymes have also been described from this organism (Morag et al., 1990). These include two free enzymes (one of which lacks a CBM) and two cell-associated (SLH-containing) enzymes. Consequently, the potent cellulose- and plant cell wall-degrading activities of C. thermocellum are clearly reflected in its cellulase system, which displays an exceptional wealth, diversity and intricacy of enzymatic components, thus representing the premier cellulose-degrading organism currently known. Many of the C. thermocellum cellulosomal enzymes are cellulases, which include both endo- and exo-acting b-glucanases. Some of the important exoglucanases and processive cellulases include CelS, CbhA, CelK and CelF. The CelS subunit is a member of the family-48 glycosyl hydrolases, and this particular family is now recognized as a critical component of bacterial cellulosomes (Morag et al., 1991; Morag et al., 1993; Wang et al., 1993; Wang et al., 1994; Wu et al., 1988). Several other processive cellulases are members of the family-9 glycosyl hydrolases. CelF and CelN are theme-B family-9 enzymes (Navarro et al., 1991; Fig. 7). The other two are remarkably similar theme-D enzymes, which exhibit nearly 95% similarity along their common regions (Kataeva et al., 1999a; Kataeva et al., 1999b; Zverlov et al., 1998; Zverlov et al., 1999). The main difference between CbhA and CelK is the presence in the former of three extra modules (a family-3 CBD and two modules of unknown function). The functional significance of these supplementary modules to the activity of CbhA has not been elucidated. The fact that the cellulosome from this organism contains many different types of cellulases is, of course, to be expected if we consider that growth of C. thermocellum is restricted to cellulose and its breakdown products, particularly cellobiose. Consequently, it is surprising to discover, in addition to the cellulases, at least five classic xylanases, i.e., those belonging to glycosyl hydrolase families 10 and 11. In addition, two of the larger enzymes, CelH and CelJ, contain hemicellulase components, i.e., family-26 and -44 cata-

CHAPTER 1.19

lytic modules (a mannanase and a xylanase, respectively), together with a standard cellulase module in the same polypeptide chain (Ahsan et al., 1996; Yagüe et al., 1990). It is also interesting to note the presence of carbohydrate esterases together with xylanase or cellulase modules in some of the enzyme subunits (i.e., XynU/A, XynY, XynZ and CelE), thus conferring the capacity to hydrolyze acetyl or feruloyl groups from hemicellulose substrates (Blum et al., 2000; Fernandes et al., 1999). Finally, the C. thermocellum cellulosome includes a typical family-16 lichenase, a family-26 mannanase and a family18 chitinase. The non-cellulosomal enzymes include another theme-B family-9 cellulase (CelI), and cell-bound forms of a xylanase (XynX) and a lichenase (LicA), both of which contain multiple CBMs adjacent to the catalytic module. In the midst of all this complexity, the C. thermocellum non-cellulosomal cellulase system includes a simple family-5 cellulase, CelC, which is completely devoid of additional accessory modules. Why does this bacterium—which subsists exclusively on cellulosic substrates—need all these hemicellulases? The inclusion of such an impressive array of non-cellulolytic enzymes in a strict cellulose-utilizing species would suggest that their major purpose would be to collectively purge the unwanted polysaccharides from the milieu and to expose the preferred substrate—cellulose. The ferulic acid esterases, in concert with the xylanase components of the parent enzymes, could grant the bacterium a relatively simple mechanism by which it could detach the lignin component from the cellulosehemicellulose composite. The lichenase (LicB) and chitinase (ChiA) are also intriguing components of the cellulosome. The former would provide the bacterium with added action on cell-wall b-glucan components from certain types of plant matter. It is not clear whether the presence of the latter cellulosomal enzyme would reflect chitin-derived substrates from the exoskeletons of insects and/or from fungal cell walls. Whatever the source, the chitin breakdown products, like those of the hemicelluloses, would presumably not be utilized by the bacterium itself, but would be passed on to appropriate satellite bacteria for subsequent assimilation.

Phylogenetics of Cellulase and Cellulosomal Systems Early in the history of the development and establishment of the cellulosome concept, it was noted that the apparent occurrence of cellulo-

CHAPTER 1.19

Cellulose-Decomposing Bacteria and Their Enzyme Systems

609

Known Enzymes from Clostridium Thermocellum Cellulosomal Enzymes X7

9

Ig

CelJ

X

44

3

48

CelS

CBM3-GH5-Doc

X7-Ig-GH9-GH44-Doc-X

CelC

GH48-Doc

CelB 9 4

X1 X1

Ig

CbhA

3

GH5 3c

GH5-Doc

GH9-CBM3c-Doc

CelN 22

10

5

CelG

9

XynY

5

5

9

CelF

CBM4-Ig-GH9-2(X1)-CBM3-Doc

Non-Cellulosomal Enzymes

5

CelO

3c

3c

3

GH9-CBM3c-CBM3

GH5-Doc

22

9

CelI

GH9-CBM3c-Doc CBM22-GH10-CBM22-Doc-CE1

XynA (XynU)

5

CelH

18

ChiA

6

9

Ig

CelA

10

X 16

LicA

8

CelD

9

9

4

4

4

4

SLH-X-GH16-4(CBM4)

9

GH8-Doc

Ig-GH9-Doc

Ig

4

22

CBM22-GH10-2(CBM9)-SLH

GH18-Doc

GH26-GH5-CBM11-Doc

CelK

XynX

GH11-CBM6-Doc-CE4

11

26

11

CBM4-Ig-GH9-Doc

22

XynC XynZ

6

10

10

XynB (XynV)

CBM22-GH10-Doc

11

6

GH11-CBM6-Doc

CEI-CBM6-Doc-GH10

CelE

ManA

5

x4

26

LicB

CBMx4-GH9-Doc

16 GH16-Doc

GH5-Doc-CE2

Scaffoldin Anchoring Proteins 3a 2Coh-CBM3a-7Coh-X-Doc

Fig. 22. Clostridium thermocellum: A very complex cellulosomal system. The modular content of the enzymes is shown from (left to right) the N-terminus to the C-terminus of the polypeptide chain. Key to symbols: GH, glycosyl hydrolase (e.g., cellulase, xylanase and mannanase); CE, carbohydrate esterase (e.g., acetyl xylan esterase and ferulic acid esterase); CBM, carbohydrate-binding module (e.g., CBD, cellulose-binding domain); Doc, dockerin domain; SLH, S-layer homology (domain); Ig, immunoglobulin-like domain; and X, domain of unknown function.

somes in different microorganisms tended to cross ecological, physiological and evolutionary boundaries (Lamed et al., 1987). Initial biochemical and immunochemical evidence to this effect has been supported by the accumulated molecular biological studies. Various lines of evidence indicate that the modular enzymes that degrade plant cell wall polysaccharides have evolved from a restricted number of common ancestral sequences. Much of the information in this direction remains as a legacy, inherently encoded in the sequences of the functional domains that comprise the different enzymes. By comparing sequences of the various cellulosomal and noncellulosomal enzymes

within and among the different strains, we can gain insight into the evolutionary rationale of the multigene families that comprise the glycosyl hydrolases.

Horizontal Gene Transfer It is clear that very similar enzymes which comprise a given glycosyl hydrolase family are prevalent among a variety of different bacteria and fungi, thus indicating that they were not inherited through conventional evolutionary processes. The widespread occurrence of such conserved enzymes among phylogenetically different species argues that horizontal transfer of

610

E.A. Bayer, Y. Shoham and R. Lamed

genes has been a major process by which a given microorganism can acquire a desirable enzyme. Once such a transfer event has taken place, the newly acquired gene would then be subjected to environmental pressures of its new surroundings, i.e., the genetic and physiological constitution of the cell itself. Following such selective pressure, the sequence of the gene would be adjusted to fit the host cell.

Gene Duplication Sequence comparisons have also revealed the presence of very similar genes within a genome that may have very similar or even identical functions. One striking example is the tandem appearance of cbhA and celK genes in the chromosome of Clostridium thermocellum. Other examples are xynA and xynB also of C. thermocellum and xynA of the anaerobic fungus Neocallimastix patriciarum, which includes two very similar copies of family-11 catalytic modules within the same polypeptide chain. These examples imply a mechanism of gene duplication (Chen et al., 1998; Gilbert et al., 1992), whereby the duplicated gene can serve as a template for secondary modifications that could result in two very similar enzymes with different properties, such as substrate and product specificities. A similar process could also account for the multiplicity of other types of modules (i.e., CBDs, cohesins or helper modules) within a polypeptide chain. Comparison of the modular architectures of similar genes from different species would suggest that individual modules can undergo a duplication process. This is exemplified by the multiple copies of FN3 in CelB from Cellulomonas fimi versus the single copy of the same domain in cellulase E4 from Thermobifida fusca. But innumerable other examples are evident from the databases, whenever multiple copies of the same modular type exist in the same protein.

Domain Shuffling Another observation from the genetic composition of the glycosyl hydrolases argues for an alternative type of process, which would propagate new or modified types of enzymes. It is clear that many microbial enzyme systems contain individual hydrolases that carry very similar catalytic domains but include different types of accessory modules (Gilkes et al., 1991). An example that demonstrates this phenomenon is the observed species preference of otherwise very similar glycosyl hydrolases for a given family of crystalline cellulose-binding CBD, which is entirely independent of the type of catalytic module borne by the complete enzyme. In this

CHAPTER 1.19

context, as we have seen above, the free enzymes of some bacteria, such as Cellulomonas fimi, Pseudomonas fluorescens and Thermomonospora fusca, invariably include a family-2 CBD, irrespective of the type of catalytic domain. In contrast, those of other bacteria, e.g., Bacillus subtilis, Caldocellum saccharolyticum, Erwinia carotovora and various clostridia, appear to prefer family-3 CBDs. Moreover, the position of the CBD in the gene may be different for different genes. For example, the CBD may occur upstream or downstream from the catalytic domain; it may be positioned either internally (sandwiched between two other modules) or at one of the termini of the polypeptide chain. The same pattern is characteristic of several other kinds of modules associated with the plant cell wall hydrolases. This is particularly evident in family-9 cellulases and family-10 xylanases, where the number and types of accessory modules may vary greatly within a given species. It seems that individual domains can be transferred en bloc and incorporated independently into appropriate enzymes. Once again, the modular architectures and sequence similarities between Clostridium thermocellum cellulosomal enzyme pairs (CbhA and CelK; XynA and XynB) are particularly revealing: in both cases, following an apparent gene duplication event, one or more additional modules appear to have been incorporated into the duplicated enzyme. Taken together, the information suggests that domain shuffling is an important process by which the properties of such enzymes can be modified and extended.

Proposed Mechanisms for Acquiring Cellulase and Cellulosomal Genes Like the free enzyme systems, the phylogeny of cellulosomal components seems to have been driven by processes that include horizontal gene transfer, gene duplication and domain shuffling. In cellulolytic/hemicellulolytic ecosystems, the resident microorganisms are usually in close contact, often under difficult conditions and in competition or cooperation with one another toward a common goal: the rapid degradation of recalcitrant polysaccharides and assimilation of their breakdown products. A possible scenario for the molecular evolution of a cellulase/hemicellulase system in a prospective bacterium could involve the initial transfer of genetic material from one microbe to another in the same ecosystem. The size and type of transferred material could vary, such as a gene or part of gene (e.g., selected functional modules) or even all or part of a gene cluster. The process could then be sustained by gene duplica-

CHAPTER 1.19

Cellulose-Decomposing Bacteria and Their Enzyme Systems

tion, which would propagate the insertion of repeated modules, e.g., the multiple cohesin domains in the scaffoldins, or even smaller units, such as the linker sequences or the duplicated calcium-binding loop of the dockerin domain. Domain shuffling can account for the observed permutations in the arrangement of domains in scaffoldin subunits from different species (Fig. 16). Finally, conventional mutagenesis would then render such products more suitable for the cellular environment or for interaction with other components of the cellulase system. The available data suggest that there are no set of rules, which would, at this stage, enable us to anticipate the nature of a given cellulase system from a given microorganism. It seems that phylogenetically dissimilar organisms can possess similar types of cellulosomal or non-cellulosomal enzyme systems, whereas phylogenetically related organisms that inhabit similar niches may be characterized by different types of enzyme systems. It is clear that to shed further light on this apparent enigma, we require more information about more types of enzyme systems. In addition to more sequences and structures, we will need more information—biochemical, physiological and ecological—to sharpen existing notions regarding the enzymatic degradation of plant cell wall polysaccharides or to formulate new ones. Acknowledgments. Grants from the Israel Science Foundation (administered by the Israel Academy of Sciences and Humanities, Jerusalem) and the Minerva Foundation (Germany) are sincerely appreciated.

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Prokaryotes (2006) 2:618–634 DOI: 10.1007/0-387-30742-7_20

CHAPTER 1.20 c i boreA

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Aerobic Methylotrophic Prokaryotes MARY E. LIDSTROM

Introduction Methylotrophic bacteria are those organisms with the ability to utilize (as their sole source of carbon and energy) reduced carbon substrates with no carbon-carbon bonds. By this definition the group includes bacteria that can grow on substrates such as methane, methanol, methylated amines, halogenated methanes and methylated sulfur species. Methylotrophic bacteria are quite widespread in nature, being found in a variety of aquatic and terrestrial habitats (King, 1992). They appear to play an important role in the cycling of carbon in specific habitats (King, 1992), and they comprise the principal biological sink for methane and other methylated greenhouse gases, highlighting an important role in global warming (King, 1992; Oremland and Culbertson, 1992). Although many anaerobic methylotrophic bacteria are known, especially among the methanogens, this chapter will cover only the aerobic and facultatively anaerobic methylotrophs (for convenience, termed “aerobic methylotrophs”). Table 1 lists the major groups of aerobic methylotrophs with examples of the genera that have been described to contain methylotrophs. Aerobic methylotrophic bacteria are phylogenetically diverse, with representatives found among the Proteobacteria as well as the high and low G+C Gram-positive bacteria (Firmicutes; Table 1). Many of the known strains of methylotrophic bacteria are obligately methylotrophic species, that is, they are incapable of growing on any compounds containing carbon-carbon bonds. However, especially among the group of bacteria that grow on methanol, a variety of facultative organisms are known that can grow either on multi-carbon compounds or on onecarbon (C1) compounds (Table 1). Two functional groups of methylotrophs may be distinguished: those capable of growth on methane, called “methanotrophs,” and those capable of growth on methanol and/or other methylated compounds but not on methane. The methanotrophs are characterized by the presence of internal membrane systems (Hanson and

Hanson, 1996). Many but not all of the methylotrophs also can use N2 as a nitrogen source and therefore are considered to be diazotrophs (Table 1). In addition, several of the methylotrophs also affect the nitrogen cycle by carrying out transformations of ammonia and nitrate (Anthony, 1982). Some methylotrophs are known that can use methylated sulfur species, and these appear to play an important role in sulfur cycling (DeBont et al., 1981; Kelly and Murrell, 1999). A number of methylotrophs can grow on halogenated methanes (Leisinger and Braus-Stromeyer, 1995) and have the potential to play an important role in the detoxification of these pollutants. The ability to grow on reduced C1 compounds requires the presence of unique biochemical pathways for both energy and carbon metabolism. So far, a limited number of variations of these metabolic pathways are known. Figure 1 gives an outline of methylotrophic metabolism, showing how different methylotrophic substrates are fed into central metabolic pathways. A key feature of aerobic methylotrophy is the role of formaldehyde as a central intermediate. In most methylotrophs, the pool of formaldehyde generated from methylotrophic substrates is split, with part being oxidized to CO2 for energy and part being assimilated into cell carbon via one of two unique pathways, the serine cycle or the ribulose monophosphate cycle. Other methylotrophs, sometimes called “pseudomethylotrophs” or “autotrophic methylotrophs” (Anthony, 1982), are capable of growth on reduced C1 compounds by oxidizing these compounds to CO2 and then assimilating the CO2 via the classical CalvinBenson-Bassham cycle. The diagram shown in Fig. 1 is an amalgam of the known diversity of methylotrophic metabolism, and no single methylotroph can carry out all of these types of metabolism. In fact, the major phylogenetic divisions mirror distinct physiological classes. For instance, all of the known methylotrophs containing the serine cycle for formaldehyde assimilation are clustered in the a-Proteobacteria, all of the restricted obligate methylotrophs that do not use methane are clustered in the b-

CHAPTER 1.20

Aerobic Methylotrophic Prokaryotes

619

Table 1. Characteristics of aerobic methylotrophic bacteria. Major assimilation pathway

N2 fixing

Obligate methylotrophs Type I methanotrophs Methylomonas Methylobacter Methylococcus Methylomicrobium Methylosphaera Methylocaldum

RuMP RuMP RuMP RuMP RuMP RuMP

Yes Yes Yes No No No

g-Proteobacteria g-Proteobacteria g-Proteobacteria g-Proteobacteria g-Proteobacteria g-Proteobacteria

Anthony, 1982 Anthony, 1982 Anthony, 1982 Bowman et al., 1995 Bowman et al., 1997 Bodrossy et al., 1997

Type II methanotrophs Methylosinus Methylocystis Methylocella

Serine Serine Serine

Yes Yes Yes

a-Proteobacteria a-Proteobacteria a-Proteobacteria

Anthony, 1982 Anthony, 1982 Dedysh et al., 2000

Restricted facultative methylotrophs Methanol utilizers Hyphomicrobium

Serine

No

a-Proteobacteria

Methylophilus

RuMP

No

b-Proteobacteria

Methylobacillus Methylophaga

RuMP RuMP

No No

b-Proteobacteria g-Proteobacteria

Harder and Attwood, 1978; Stackebrandt et al., 1988 Jenkins and Jones, 1987 Bratina et al., 1992 Janvier and Grimont, 1995

Serine

No

a-Proteobacteria

Aminobacter Methylorhabdus Methylopila Methylosulfonomonas Marinosulfonomonas Paracoccus Xanthobacter Ancylobacter (Microcyclus) Thiobacillus

Serine Serine Serine Serine Serine CBB CBB CBB CBB

No No No No No No Yes Yes No

a-Proteobacteria a-Proteobacteria a-Proteobacteria a-Proteobacteria a-Proteobacteria a-Proteobacteria a-Proteobacteria a-Proteobacteria a-Proteobacteria

Rhodopseudomonas Rhodobacter Acetobacter Bacillus

CBB CBB RuMP RuMP

No No ND ND

a-Proteobacteria a-Proteobacteria g-Proteobacteria Gram-positive (low G+C)

Mycobacterium

RuMP

ND

Gram-positive (high G+C)

Arthrobacter Amycolatopsis (Nocardia)

RuMP RuMP

ND ND

Gram-positive (high G+C) Gram-positive (high G+C)

Group

Facultative methylotrophs Methylobacterium (Pseudomonas)

Phylogenetic positiona

References

Green and Bousfield, 1983 Urakami et al., 1992 Doronina et al., 1995 Doronina et al., 1998 Holmes et al., 1997 Holmes et al., 1997 Anthony, 1982 Jenni et al., 1987 Raj, 1989 Chandra and Shethna, 1977 Anthony, 1982 Anthony, 1982 Yamada et al., 1997 Dijkhuizen et al., 1988 Reed and Dugan, 1987 Levering et al., 1981 De Boer et al., 1990

Abbreviations: RuMP, ribulose monophosphate; CBB, Calvin-Benson-Bassham; and ND, no data. Many phylogenetic affiliations can be found at the Ribosomal Database Project (http://www.cme.msu.edu/RDP/ html/index.html) or National Center for Biotechnology Information websites and in Bratina et al. (1992).

a

Proteobacteria (with one exception, Methylophaga), all of the methanotrophs that use the ribulose monophosphate cycle for formaldehyde assimilation are clustered in the g-Proteobacteria, and all of the known Gram-positive methylotrophs contain the ribulose monophosphate cycle (Table 1). For the methanotrophs, the a-

Proteobacteria containing the serine cycle are referred to as type I strains, whereas the gProteobacteria containing the ribulose monophosphate cycle are referred to as type II strains (Hanson and Hanson, 1996). Because the natural diversity of methylotrophs is still under investigation, the current clustering of phylogenetic

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M.E. Lidstrom

O2 CH4 2H

1

CH3OH

CHAPTER 1.20

Methylated amines Methylated sulfur species O2 6 7 NH3 2 5 S NH3 H2O2 H2 O 4 3 2 HCOOH CO2 HCHO 2H

H 2O

RuMP Cycle

2H

Serine Cycle

2H

RuBP Cycle

3-Carbon compounds Cell Material

Fig. 1. Metabolism of one-carbon compounds in aerobic methylotrophic bacteria. 1, methane monooxygenase; 2, methanol dehydrogenase; 3, formaldehyde oxidation system; 4, formate dehydrogenase; 5, halomethane oxidation system; 6, methylated amine oxidases; 7, methylated amine dehydrogenase; and 8, methylated sulfur dehydrogenase or oxidase. RuMP is ribulose monophosphate, and CBB is CalvinBenson-Bassham. Adapted from Anthony (Anthony, 1982; Anthony, 1996) and DeBont et al. (1981).

and physiological groups may not hold up, as new strains are identified and characterized.

Dissimilatory Metabolism Aerobic methylotrophs contain specialized pathways for dissimilatory metabolism during methylotrophic growth. In general, the methyl groups of methylotrophic substrates are oxidized to the level of formaldehyde by oxidases and/or dehydrogenases. The dehydrogenases are generally coupled to energy metabolism at the level of cytochromes and the oxidases are usually nonenergy conserving. Formaldehyde is then further oxidized to the formyl level by one of a number of formaldehyde oxidation systems, which usually generate a reduced pyridine nucleotide. Carbon at the level of formate is then oxidized to CO2 via another pyridine nucleotide-linked step.

Methane Oxidation The enzyme that oxidizes methane to methanol in the methanotrophic bacteria is a mixedfunction oxidase called “methane monooxygenase” (MMO; Fig. 1). Two different enzymes are known, a membrane-bound form, known as “the particulate MMO” (pMMO), and a soluble form, called “the soluble MMO” (sMMO; Hanson and Hanson, 1996). The soluble MMO has so far been documented in only a few strains and it is not yet known how widely distributed it is in methanotrophs. It has been found in all tested strains of Methylosinus and Methylococcus, and in a few strains of Methylomonas Methylomicro-

bium and Methylocystis (Hanson and Hanson, 1996; Fuse et al., 1998; Shigematsu et al., 1999; Grosse, 1999). However, pMMO appears to be present in all known strains of methanotrophs. The sMMO has been purified from both type I and type II methanotrophs (Lipscomb, 1994), and it is similar in all cases. It consists of three components: a hydroxylase (consisting of three polypeptides and a non-heme iron center), component B (with no cofactors), and a reductase that contains FAD and and an Fe2S2 cluster (Lipscomb, 1994). The sMMO uses NADH as a source of reducing power, and contains an hydroxo-bridged di-iron center in its active site (Lipscomb, 1994). It is characterized by an extremely broad substrate specificity, being able to oxidize or hydroxylate a wide variety of aliphatic straight chain, branched, aromatic, and halogenated hydrocarbons (Lipscomb, 1994; Hanson and Hanson, 1996). The broad substrate range of this enzyme has attracted a great deal of attention as a result of the use of methanotrophs for bioremediation of a variety of toxic hydrocarbons (Hanson and Hanson, 1996). Crystal structures are available for the hydroxylase and component B from two different methanotrophs (Walters et al., 1999; Elango et al., 1997; Chang et al., 1999; Rosenzweig et al., 1997). Genes for the subunits of the sMMO (mmo genes) have been cloned and sequenced from a number of methanotrophs, and they have a similar organization with high similarity at the amino acid level (Murrell, 1994; Shigematsu et al., 1999; McDonald et al., 1997). The pMMO is highly unstable and has proven more difficult to analyze. However, recently, pMMO was purified in an active state from Methylococcus capsulatus Bath by two groups (Zahn and DiSpirito, 1996; Nguyen et al., 1998). In both cases, the pMMO had 3 subunits, PmoABC, of approximately 27, 45 and 22 kDa, respectively, and was a copper-containing enzyme. In one case no other metals were present (Nguyen et al., 1998), whereas in the other iron was also present (Zahn and DiSpirito, 1996). The pMMO has a narrower substrate range than the sMMO (Hanson and Hanson, 1996). The genes encoding the pMMO are present in multiple copies in most methanotrophs (Semrau et al., 1995). In both a type I and type II methanotroph, evidence exists that the copies are nearly identical in sequence (Stolyar et al., 1999; Gilbert et al., 2000), and mutant evidence has shown that in Methylococcus capsulatus Bath the copies are functionally redundant (Stolyar et al., 1999). In methanotrophs containing both pMMO and sMMO, the expression of each enzyme is regulated by copper. In copper sufficiency, pMMO is expressed, and in copper limitation,

CHAPTER 1.20

sMMO is expressed. In Methylococcus capsulatus Bath and Methylosinus trichosporium, it has been shown that this regulation occurs at the transcriptional level (Nielsen et al., 1997).

Methanol Oxidation Methanol is widespread, produced in nature as a result of demethylation reactions (Anthony, 1982), especially from plants (Holland and Polacco, 1994). Methanol is oxidized to formaldehyde by three classes of enzymes, a quinoprotein methanol dehydrogenase (MDH) found in the Gram-negative methylotrophs (Goodwin and Anthony, 1998), an NAD-linked enzyme found in the Bacillus strains (Arfman et al., 1997), and a methanol:N,N¢-dimethyl-4nitrosoaniline oxidoreductase (MNO) found in other Gram-positive strains (Bystrykh et al., 1993; Bystrykh et al., 1997). In general, methanol oxidation is an energy-conserving step, either generating reduced cytochromes or reduced pyridine nucleotides.

Quinoprotein Methanol Dehydrogenase All of the known Gram-negative methanol- and methane-utilizing bacteria contain a periplasmic enzyme for oxidizing methanol called “methanol dehydrogenase.” This enzyme, which oxidizes primary alcohols to their corresponding aldehydes, has an a2b2 structure and contains the cofactor pyrroloquinoline quinone (PQQ; Goodwin and Anthony, 1998). Electrons from the oxidation of PQQ are transferred from PQQ to a specific cytochrome c, and from there through other carriers to the terminal oxidase (Goodwin and Anthony, 1998). The primary sequences and structures for methanol dehydrogenase from diverse methylotrophs are highly conserved (Goodwin and Anthony, 1998). These enzymes contain a Ca2+ near the active site and also have an unusual disulfide bridge in the same region (Goodwin and Anthony, 1998; Anthony and Ghosh, 1998).

NAD-Linked Methanol Dehydrogenases An NAD-linked methanol dehydrogenase has been purified and characterized from methylotrophic Bacillus strains (Arfman et al., 1997). This enzyme oxidizes C1–C4 primary alcohols, and is composed of ten identical 43,000-Mr subunits. Each MDH subunit contains a tightly, but noncovalently bound NAD(H) molecule, in addition to 1 Zn2+ and 1 or 2 Mg2+ ions. This MDH also interacts with a 50,000-Mr activator protein, which appears to facilitate the oxidation of the reduced NADH cofactor of MDH (Arfman et al., 1997). The structural gene for

Aerobic Methylotrophic Prokaryotes

621

this MDH shows identity with type II alcohol dehydrogenases (de Vries et al., 1992).

Methanol:N,N¢-dimethyl-4-nitrosoaniline Oxidoreductase (MNO) Other Gram-positive methylotrophs (Amycolatopsis and Mycobacterium) oxidize methanol via a methanol:N,N¢-dimethyl-4-nitrosoaniline oxidoreductase (MNO), which is a decameric protein with 50-kDa subunits, each carrying a tightly bound NADPH (Bystrykh et al., 1997). This protein also has been isolated as a complex containing two other components that impart a tetrazolium-dye-linked methanol dehydrogenase activity (Bystrykh et al., 1997).

Oxidation of Methylated Amines Methylated amines are also widespread in the environment, being produced as degradation products of some pesticides, of carnitine and lecithin derivatives, and of trimethylamine oxide. The latter is especially prevalent in fish and in marine environments (Anthony, 1982). A variety of bacteria are known that are capable of growing on methylated amines. In general, the methyl groups of methylated amines are oxidized to formaldehyde, either by an oxidase or a dehydrogenase, with energy conservation occurring in the latter case. Growth on formaldehyde occurs via normal methylotrophic assimilatory and dissimilatory pathways (Fig. 1).

Trimethylamine and Dimethylamine Trimethylamine is oxidized to dimethylamine and formaldehyde by trimethylamine dehydrogenase (Fig. 2). This enzyme is a flavoprotein, that also contains two Fe2S2 clusters and two molecules of ADP (McIntire, 1990). A second pathway for utilization of trimethylamine occurs in which a trimethylamine monooxygenase oxidizes trimethylamine to trimethylamine N-oxide. The N-oxide is subsequently demethylated by trimethylamine demethylase to dimethylamine and formaldehyde (Anthony, 1982). Dimethylamine is oxidized to methylamine and formaldehyde by dimethylamine monooxygenase (Fig. 2). Gene sequences suggest that trimethylamine and dimethylamine dehydrogenases are evolutionarily related (Yang et al., 1995).

Methylamine Four possible routes are known in bacteria for utilizing methylamine (Fig. 2). The first of these involves the periplasmic enzyme, methylamine dehydrogenase (MADH), which is another quinoprotein shown to contain the cofactor,

622

M.E. Lidstrom

CHAPTER 1.20 2H

H2O Trimethylamine

1

(CH3)3N O2

O2, NADH Dimethylamine

CH3NH2 NH3

NADPH HCHO

NH3

5

NADP

Trimethylamine N-Oxide (CH3)3NO

Methylamine

4

(CH3)2NH

2 H2O

H2O, NAD

2H

3

O2

8

H2O2

6 7 9

glutamate

g -Glutamylmethylamide

N-methylglutamate

2H H2 O

Fig. 2. Pathways for converting methylated amines to formaldehyde in methylotrophic bacteria. 1, trimethylamine dehydrogenase; 2, trimethylamine monooxygenase; 3, trimethylamine N-oxide demethylase; 4, dimethylamine monooxygenase; 5, methylamine dehydrogenase; 6, methylamine oxidase; 7, N-methylglutamate synthase; 8, g-glutamylmethylamide synthetase; and 9, N-methyl glutatmate dehydrogenase. From Anthony (1982).

tryptophan tryptophylquinone (TTQ), instead of the PQQ found in MDH. This TTQ is formed by covalent crosslinking of two tryptophan residues in the small subunit of MADH, and incorporation of two oxygen atoms into one of the indole rings to form a quinone (Davidson, 1999). The MADH converts primary amines to their corresponding aldehydes plus ammonia, and electrons are transferred to a small copper protein, amicyanin. These electrons are transferred to the respiratory chain via a c-type cytochrome (Davidson, 1999). Structural, kinetic and sitedirected mutagenesis studies have characterized protein-protein interactions, and mechanisms of catalysis and electron transfer by TTQ. In addition, the genes encoding the functions required for active MADH (mau genes) have been studied from several bacteria, and they are similar in both amino acid sequence and genetic organization (van der Palen et al., 1995; Chistoserdov, 1994a; Chistoserdov, 1994b; Gak et al., 1997; Graichen et al., 1999). In Arthrobacter P1, methylamine is utilized by another quinoprotein, methylamine oxidase. This enzyme is a blue copper amine oxidase similar to mammalian copper amine oxidases, which generate hydrogen peroxide (Levering et al., 1981; McIntire and Hartman, 1993; Fig. 2). This enzyme contains the cofactor 6-hydroxydopa quinone, which is formed posttranslationally from a tyrosine residue in the amino acyl chain (McIntire and Hartman, 1993). A few methylotrophs contain indirect pathways, which involve the conversion of methylamine to Nmethylglutamate, and finally to formaldehyde; N-methylglutamate can be synthesized directly, via N-methylglutamate synthase, or indirectly via a g-glutamylmethylamide intermediate, as shown in Fig. 2, although the latter pathway is still uncertain (Anthony, 1982).

Utilization of Methylated Sulfur Species A few organisms have been isolated that are capable of utilizing methylated sulfur compounds such as dimethylsulfoxide (DMSO), dimethylsulfide (DMS), and dimethyldisulfide (DMDS). Most of these strains have been Hyphomicrobium species (DeBont et al., 1981; Suylen and Kuenen, 1986), but a few Thiobacillus strains and a Methylophaga strain have been reported (Kanagawa and Kelly, 1986; De Zwart et al., 1996). The Hyphomicrobium strains apparently reduce DMSO to DMS and then convert the DMS to methanethiol and formaldehyde (Fig. 3). The methanethiol is then oxidized by an oxidase to H2S and formaldehyde with the production of hydrogen peroxide (DeBont et al., 1981). Formaldehyde is utilized by standard methylotrophic pathways (Fig. 1). DMS arises in marine environments through the cleavage of dimethyl-b-propiothetin, one of the products of sulfur metabolism of marine algae (Andreae, 1980). Also, DMS has been thought to play an important role in the transport of reduced sulfur compounds between aquatic and terrestrial environments and from terrestrial environments into the atmosphere (Andreae and Raemdonck, 1983; Banwart and Bremner, 1976). The distribution of bacteria capable of utilizing methylated sulfur compounds has not been well studied. However, it seems likely that these organisms are widespread and are present in many environments in which DMS is produced. Some specialized methylotrophs (including Methylosulfonomonas, Marinosulfonomonas, and strains of Hyphomicrobium and Methylobacterium) can use methanesulfonate as a carbon and energy substrate to support growth (Kelly and Murrell, 1999; Pol et al., 1994). Methane-

CHAPTER 1.20 Fig. 3. Proposed pathway for converting methylated sulfur species to formaldehyde in Hyphomicrobium species Adapted from DeBont et al. (1981).

Aerobic Methylotrophic Prokaryotes Methylated amines

Halomethanes

Methylated sulfur species (O2)

O2

8

7 5

6 NH3

X–, 2H O2

1

CH4 2H

CH3OH

S=

NH3

H2O2 H2O

2

3

HCHO 2H

H2O

623

2H

4 HCOOH or Formyl 2H Derivatives

CO2

α-proteobacteria

β and γ-proteobacteria

RuMP Cycle

CBB Cycle

Serine Cycle α-proteobacteria

3-Carbon compounds Cell Material

sulfonate is oxidized to sulfite and formaldehyde by NADH-dependent methanesulfonate monooxygenase, and utilization of formaldehyde proceeds by normal serine-cycle-dependent methylotrophic metabolism (Kelly and Murrell, 1999). The methanesulfonate monooxygenase has been shown to consist of three components: 1) a 200-kDa hydroxylase complex containing two major polypeptides of around 50 and 20 kDa with a Rieske [2Fe-2S] center; 2) a 16-kDa ferredoxin component; and 3) the putative reductase component, a 36-38 kDa-monomeric protein catalyzing the NADH-dependent reduction of several electron acceptors, including cytochrome c (Kelly and Murrell, 1999).

Halomethanes A number of methylotrophic bacteria are known that are capable of aerobic growth on halomethanes such as chloromethane, bromomethane and dimethylchloride (Leisinger, 1994; Leisinger and Baus-Stromeyer, 1995; Hancock et al., 1998). These bacteria are generally found in the genera Methylobacterium, Hyphomicrobium or Methylophilus, although two strains using monohalomethanes also have been identified that class together in a new subgroup of a-proteobacterial methylotrophs within a clade of rhizobia (Schaefer, 1999; Coulter, 1999). Dichloromethane degradation involves a glutathionelinked dehalogenase that produces formaldehyde (Leisinger, 1994), and the rest of metabolism proceeds by general methylotrophic pathways. Chloromethane degradation has been shown to involve a corrinoid-dependent methyltransferase with sequence identity to methanogen methyltransferases (Studer et al., 1999; Coulter, 1999). In Methylobacterium strain CM4,

the methyltransferase reaction is coupled to tetrahydrofolate derivatives to produce formate, followed by formate oxidation (Vannelli, 1999; Fig. 4). Assimilation occurs via methylene tetrahydrofolate and the serine cycle. In strain CC495, which is one of the chloromethane utilizers that classes near rhizobia, evidence is presented for a bisulfide-coupled reaction in which methanethiol is the product (Coulter, 1999). In that case, it has been proposed that methanethiol is oxidized to formaldehyde, and metabolism proceeds by general methylotrophic pathways.

Formaldehyde Oxidation Although it is theoretically possible for methylotrophs to grow on formaldehyde, this substrate is usually too toxic to sustain growth in batch cultures. A few cultures of both methane and methanol utilizers have been reported to grow on formaldehyde (Whittenbury et al., 1981; Hirt et al., 1978), but the growth is poor and usually requires that the substrate be provided in the gas phase. Arthrobacter P1 has been grown in a formaldehyde-limited chemostat by first establishing cultures on choline, then adding low levels of formaldehyde, and finally eliminating the choline gradually (Levering et al., 1986). It seems likely that other methylotrophs could be grown on formaldehyde using a similar technique. A number of formaldehyde oxidation systems are known in methylotrophs (Figs. 4 and 5). The simplest of these is formaldehyde dehydrogenase, which converts formaldehyde to formate. A number of NAD-linked and dye-linked (presumably PQQ-containing and cytochromelinked) formaldehyde dehydrogenases have been identified from methylotrophs, but the low activity and general lack of inducibility of these

624

M.E. Lidstrom

CHAPTER 1.20

NAD NADH2 A. Formaldehyde

Formate NAD-dependent FaDH GSH

NAD NADH2

GSH

S-hydroxy methyl GSH

B. Formaldehyde

S-formyl GSH

NAD-GSH dependent FaDH

(spontaneous)

Fomate S-formyl GSH hydrolase

MySH

NAD NADH2

MySH C. Formaldehyde (spontaneous)

S-hydroxy methyl MySH

S-formyl MySH

Formate

NAD-MySH dependent FaDH

NAD(P) NAD(P)H2

2e-, MFR?

MFR? H4MPT N 5-Formyl

Methenyl

CO2 Formyl MFR ? H MPT H MPT 4 4 H4MPT Methylene H4MPT (Formyl MFR Methenyl H4MPT Formyl MF R:H4MPT dehydrogenase dehydrogenase) cyclohydrolase formyltransferase

Methylene H4MPT D. Formaldehyde

NADP NADPH

H4F

Methylene H4F

ATP, H4F

Methenyl H4F

Methylene H4F dehydrogenase

N 10-Formyl

NAD

Formate

H4F Methenyl H4F Cyclohydrolase

Formyl H4F synthetase

NADH CO2

Formate dehydrogenase

Fig. 4. Linear pathways for formaldehyde oxidation in aerobic methylotrophic bacteria. A, NAD-linked formaldehyde dehydrogenase (FaDH); B, glutathione (GSH)-linked FaDH; C, Mycothiol (MySH)-linked FaDH; D, the two formate-linked pathways, one (upper) involving tetrahydromethanopterin (H4MPT) and the other (lower) involving tetrahydrofolate (H4F). Adapted from Anthony (1982); Misset-Smiths et al. (1997); Harms (1996); Chistoserdova et al. (1998). HCHO

Cyclic Pathway

hexulose-6-P

ribulose–5–P – CO2

fructose–6–P

2H

glucose–6–P

Duine, 1990; Chistoserdova et al., 1991; Attwood et al., 1992; Speer et al., 1994). It is likely that these enzymes are involved in formaldehyde detoxification rather than playing a major dissimilatory role (Chistoserdova et al., 1991; Vorholt et al., 1999).

2H 6–PHOSPHOGLUCONATE DEHYDROGENASE

6–phosphogluconate

GLUCOSE–6–PHOSPHATE DEHYDROGENASE

Fig. 5. Cyclic pathway of formaldehyde oxidation, involving enzymes of the RuMP pathway. Adapted from Anthony (1982).

enzymes has called their physiological role into question (Hirt et al., 1978; Stirling, 1978; Anthony, 1982; Marison and Attwood, 1982; Weaver and Lidstrom, 1985; Van Ophem and

NAD- and Mycothiol-Linked Formaldehyde Dehydrogenase Recent studies have suggested that alternate routes are involved in formaldehyde dissimilation. In Gram-positive methylotrophs, the major formaldehyde dehydrogenase appears to be an enzyme that had previously been described as NAD-linked factor-dependent formaldehyde dehydrogenase (Van Ophem et al., 1992; Van Ophem and Duine, 1994; Duine,

CHAPTER 1.20

1999; Fig. 4). It is now known that this factor is mycothiol (1-O-(2¢-[N-acetyl-L-cysteinyl]amido2¢-deoxy-a-D-glucopyranosyl)-D-myoinositol), a compound also found in Mycobacterium strains (Misset-Smiths et al., 1997; Duine, 1999). This trimeric enzyme consists of a single type of subunit containing Zn (Van Ophem et al., 1992).

NAD- and GSH-Linked Formaldehyde Dehydrogenase An analogous enzyme coupled to glutathione (GSH) is involved in formaldehyde dissimilation in a variety of Gram-negative methylotrophs, including Paracoccus and Rhodobacter (Ras et al., 1995; Harms et al., 1996; Barber and Donohue, 1998; Fig. 4). In this case, two enzymes act in concert, an NAD- and GSH-linked dehydrogenase that generates the formyl-GSH derivative, and a hydrolase that releases GSH and formate. Analysis of the genes encoding these enzymes (flh genes) suggests they are similar to genes involved in formaldehyde detoxification in a variety of organisms (Harms et al., 1996; Barber and Donohue, 1998).

Folate-Linked Formaldehyde Oxidation Pathways Two additional linear formaldehyde oxidation pathways are known in methylotrophs, both linked to folates. The first of these involves a standard tetrahydrofolate (H4F) oxidation pathway, similar to the C1 interconversion pathways found in most organisms, which oxidizes methylene tetrahydrofolate to formate and tetrahydrofolate (Fig. 4). This pathway was suggested to be the major dissimilatory route for formaldehyde oxidation in serine cycle methylotrophs (Marison and Attwood, 1982). However, in Methylobacterium extorquens AM1, this pathway appears to be a minor one (Chistoserdova et al., 1998). In addition, the methylene H4F dehydrogenase is unusual for bacteria, in that it only carries out this first step instead of both steps as does the normal, coupled enzyme (encoded by folD). The gene sequence is highly divergent from other methylene H4F dehydrogenases (Chistoserdova and Lidstrom, 1994b), and Methylobacterium extorquens also contains an unusual methenyl H4F cyclohydrolase (Pomper et al., 1999). The major dissimilatory pathway has been suggested to be an analogous tetrahydromethanopterin-linked (H4MPT-linked) pathway (Fig. 4), similar to that found in the archeaon Archaeoglobus fulgidis, and to the reversal of the first few steps of the CO2 reduction pathway found in archaeal methanogens. This pathway

Aerobic Methylotrophic Prokaryotes

625

involves a cofactor that had been thought to be specific to archaea, tetrahydromethanopterin (H4MPT), and the genes encoding the archaeallike enzymes show significant identity to the corresponding archaeal genes (Chistoserdova et al., 1998). Therefore, it has been suggested that this pathway was acquired by an early methylotroph by horizontal gene transfer from an archaea (Chistoserdova et al., 1998). The archaeal H2- or F420- enzymes that interconvert methylene and methenyl H4MPT are not found in the aerobic methylotrophs. Instead, two enzymes have been identified that oxidize methylene H4MPT to methenyl H4MPT, one linked to NAD and the other to NADP (Vorholt et al., 1998; Hagemeier et al., 2000). The NAD-linked enzyme is specific to H4MPT and shows similarity to the NADP-linked enzyme (Hagemeier et al., 2000). The NADP-linked enzyme has activity with both H4MPT and H4F, although the activity with the latter is 10% that of the former, and it appears to be the only methylene H4Fdehydrogenase during methylotrophic growth (Vorholt et al., 1998). Activity and genes are present for methanofuran-utilizing enzymes, but no evidence exists for the presence of methanofuran in methylotrophs. Therefore, the details of the final oxidation step in this pathway are still not known.

Cyclic Formaldehyde Oxidation Pathway Another formaldehyde pathway is cyclic and involves the condensation of the C1 compound with a five-carbon acceptor molecule, followed by oxidation of the resulting six-carbon compound (Fig. 5). The enzymes carrying out these reactions are those of the ribulose monophosphate cycle for formaldehyde assimilation, with the exception of one novel enzyme, the 6-phosphogluconate dehydrogenase. A second enzyme is also needed, glucose-6-phosphate dehydrogenase, but this may or may not be a part of the RuMP cycle depending upon the variant utilized. These genes have been cloned and sequenced from Methylobacillus flagellatum (Chistoserdova et al., 2000). In those organisms that carry out the cyclic pathway of formaldehyde oxidation, glucose-6-phosphate dehydrogenase activity utilizes both NADP and NAD. However, in Methylophilus methylotrophus and Methylobacillus flagellatum, two different 6-phosphogluconate dehydrogenases have been found, one of which is active with both NADP and NAD, and the other specific for NAD only (Beardsmore et al., 1982; Kiriuchin et al., 1988). It has been speculated that the flow of carbon at the branch point between oxidation and assimilation in the cyclic pathway is regulated allosterically by these two isoenzymes (Beardsmore et al., 1982).

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Distribution of Formaldehyde Oxidation Pathways in Methylotrophs Although some methylotrophs appear to have only one dissimilatory formaldehyde oxidation pathway, others have multiple routes. Many methylotrophs contain low activities of one or more formaldehyde dehydrogenases, and these may play a largely protective role in formaldehyde detoxification. Of the main dissimilatory pathways, the mycothiol-linked formaldehyde dehydrogenase has so far been found only in Gram-positive methylotrophs (Duine, 1999). The GSH-linked formaldehyde oxidation system has been found mainly in the Gram-negative autotrophic methylotrophs (Harms, 1996; Barber and Donohue, 1998), whereas the H4Flinked pathway is found in the serine cycle methylotrophs (Vorholt et al., 1999). The cyclic formaldehyde oxidation pathway occurs mainly in the obligate methylotrophs containing the RuMP cycle (Anthony, 1982; Grundig and Babel, 1987). Although it also is found in Arthrobacter P1 (Levering et al., 1981), it has been found not to be effective in formaldehyde resistance and may not be a major dissimilatory pathway (Grundig and Babel, 1989). Likewise, although enzyme activities of both pathways exist in the obligate methanotrophs containing the RuMP cycle, the low activities of the glucose6-phosphate and 6-phosphogluconate dehydrogenases suggest that an alternate pathway must dominate in vivo (Zatman, 1981). This latter conclusion is supported by the fact that the H4MPTlinked pathway is found in both serine-cycle and RuMP-cycle methanotrophs, at high activity (Vorholt et al., 1999). This pathway has a broad distribution, being found in all tested Gramnegative methylotrophs with either the serine cycle or the RuMP cycle (Vorholt et al., 1999). It was not found in the Gram-positive methylotrophs tested, nor in most of the autotrophic methylotrophs. However, it was present in the autotrophic Xanthobacter strains (Vorholt et al., 1999). These results suggest that methylotrophs either have one of the thiol-linked formaldehyde oxidation systems (mainly Gram-positive and autotrophic methylotrophs), or they have the H4MPT-linked formaldehyde oxidation system (all other methylotrophs). In addition, they may have the H4F-linked pathway or the cyclic oxidation pathway. In the b-proteobacterial obligateRuMP-cycle methylotroph Methylobacillus flagellatum KT, mutational analysis suggested that the cyclic oxidation pathway was the major dissimilatory pathway, whereas the H4MPTlinked pathway played a detoxification role (Chistoserdova et al., 2000). An analysis of partial sequences of genes encoding one of the diagnostic enzymes for the H4MPT-linked pathway,

CHAPTER 1.20

methenyl H4MPT cyclohydrolase, showed that the bacterial genes grouped in a cluster separate from the cluster of archaeal genes, and the branching pattern for the bacterial genes roughly mirrored the branching pattern of the 16S rRNA genes (Vorholt et al., 1999).

Formate Oxidation In methylotrophs that have one of the linear oxidation pathways, formate is generally thought to be oxidized to CO2 by an NAD-linked formate dehydrogenase (Anthony, 1982). Two classes of soluble formate dehydrogenase have been identified in methylotrophs, a dimeric enzyme described in a Mycobacterium strain (Galkin, 1995) and in an unidentified Gram-negative methylotroph (Lamzin et al., 1992); these show similar properties and gene sequences (Galkin et al., 1995). However, most methylotrophs appear to have an enzyme (composed of 4 subunits, containing iron, molybdenum and flavin; Jollie and Lipscomb, 1991) similar in properties and at the gene sequence level to a formate dehydrogenase found in Ralstonia eutropha (Friedebold and Bowien, 1993). A membrane-bound formate dehydrogenase has been reported in Amycolatopsis methanolica, but it has not been characterized (Khmelenina et al., 1997). No mutants have been reported in formate dehydrogenase in a methylotroph, and so the physiological role of this enzyme has not yet been confirmed.

Assimilatory Metabolism Three main pathways of assimilatory metabolism are known in aerobic methylotrophs: two that assimilate carbon at the level of formaldehyde (the serine cycle and the ribulose monophosphate cycle), and one that assimilates carbon at the level of CO2 (the Calvin-Benson-Bassham or CBB cycle). Both pathways for the assimilation of formaldehyde involve cycles in which a condensation reaction between a C1 compound and a multicarbon compound occurs, followed by regeneration of the acceptor molecule and production of a C3 compound (Figs. 1, 6 and 7).

Serine Cycle The serine cycle for formaldehyde assimilation is shown in Fig. 6. This pathway initiates with the condensation of methylene tetrahydrofolate and glycine to form serine. This 3-carbon compound then undergoes a series of transformations to phosphoenolpyruvate, which is carboxylated to form malate. The malate is cleaved into two 2carbon compounds, which are then converted

CHAPTER 1.20

Aerobic Methylotrophic Prokaryotes

Fig. 6. The serine cycle for formaldehyde assimilation. Adapted from Anthony (1982).

627

3-phosphoglycerate

2 2-phosphoglycerate ADP GLYCERATE KINASE

ATP

2 glycerate NAD+ HYDROXYPYRUVATE REDUCTASE NADH + H+ hydroxypyruvate SERINE 2 serine TRANSHYDROXYMETHYLASE 2 THF 2 N5,10 methylene THF 2 glycine 2 THF 2 formaldehyde

back into glycine, thus completing the cycle. In most organisms that have the serine cycle, it is not clear how acetyl CoA is converted to glyoxylate. The usual route for this conversion involves isocitrate lyase, but this enzyme is only present in a few strains (Anthony, 1982; Chistoserdova and Lidstrom, 1996). The enzymes specific to the serine cycle are noted in the figure and most of the genes encoding these enzymes have been cloned and sequenced from Methylobacterium extorquens AM1 (Chistoserdova and Lidstrom, 1994a; Chistoserdova and Lidstrom, 1994b; Chistoserdova and Lidstrom, 1996; Chistoserdova and Lidstrom, 1997) or Hyphomicrobium methylovorum (Yoshida et al., 1994; Hagishita et al., 1996; Tanaka et al., 1997). Two isoenzymes are known to exist for phosphenol pyruvate (PEP) carboxylase (Newaz and Hersh, 1975; McNerney and O’Connor, 1980). The C1-specific PEP carboxylase is acetyl-CoA independent, unlike the classical acetyl-CoA-dependent anapleurotic enzyme (Newaz and Hersh, 1975), and the gene sequence for this enzyme is on the order of 30% identical to genes encoding the anapleurotic enzyme (Chistoserdova and Lidstrom, 1997). For each C3 compound that is generated by the serine cycle two carbons are derived from formaldehyde and one from CO2.

Ribulose Monophosphate Cycle The ribulose monophosphate cycle (RuMP cycle) is shown in Fig. 7. Formaldehyde is condensed with the acceptor molecule (ribulose monophosphate) by the enzyme hexulose phosphate synthase to produce hexulose phosphate. The six-carbon molecule is then isomerized to fructose 6-phosphate by phosphohexulose isomerase, and a series of interconversions occur that regenerate the five-carbon acceptor molecule. The condensation of three formaldehyde

phosphoenolpyruvate CO2 PEP CARBOXYLASE oxaloacetate NADH + H+ SERINE-GLYOXYLATE NAD+ AMINOTRANSFERASE malate NH2 ATP MALATE ADP THIOKINASE malyl CoA MALYL CoA LYASE glyoxylate acetyl CoA glyoxylate

molecules results in the net production of one C3 compound (Anthony, 1982). As shown in Fig. 7, four different variants of the ribulose monophosphate pathway are possible. However, only three of the possible combinations have been shown to exist. In those obligate methylotrophs that use the RuMP pathway, the combination that appears to occur is that involving 6-P-gluconate (right) and not dihydroxyacetone phosphate (left). The two key enzymes for this variant are 2-keto, 3-deoxy, 6-phosphogluconate aldolase and transaldolase. In the facultative methylotrophs the existing evidence suggests that some strains such as Bacillus PM6 (Colby and Zatman, 1975) contain the combination involving fructose bisphosphate (center) and sedoheptulose bisphosphate (lower), and the two key enzymes in this case are fructose bisphosphatase and sedoheptulose bisphosphatase. Other facultative strains, such as Arthrobacter P1, contain the combination involving the enzymes fructose bisphosphatase (center) and transaldolase (left; Levering et al., 1982). Genes encoding hexulose phosphate synthase and phosphohexuloisomerase (rmpA and rmpB) have been cloned and sequenced from both Gram-positive and Gram-negative methylotrophs (Yanase et al., 1996; Sakai et al., 1999; Mitsui et al., 2000). Surprisingly, genes with identity to hexulose phosphate synthase are common in non-methylotrophic bacteria and in archaea, and for the most part their role is not known (Reizer et al., 1997). However, Bacillus subtilis contains orthologs to both rmpA and rmpB, and evidence has been presented that these genes encode functional enzymes involved in protection of the cell from formaldehyde (Yasueda et al., 1999). Those organisms that utilize the CalvinBenson-Bassham pathway for CO2 fixation appear to utilize the standard pathway without alternations (Anthony, 1982).

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CHAPTER 1.20

3 HCHO

PHOSPHOHEXULOSE ISOMERASE 3 ribulose–5–P 3 hexulose–6–P fructose–6–P HEXULOSE PHOSPHATE ATP SYNTHASE fructose–6–P fructose–6–P

fructose–6–P or

ADP

glyceraldehyde–3–p ribose–5–P

fructose1,6–bis P

glucose–6–P NADP NADPH+H+ 6–P–gluconate

erythrose–4–P sedoheptulose–7–P glyceraldehyde–3–P dihydroxyocetone–P xylose–5–P xylose–5–P

2-ketodeoxy phosphogluconate

glyceraldehyde–3–P pyruvate

or fructose–6–P ATP ADP fructose1,6–bis P

glyceraldehyde–3–P dihydroxyacetone–P erythrose–4–P sedoheptulose1,7–bis P

sedoheptulose–7–P Pi

Fig. 7. The ribulose monophosphate (RuMP) cycle for formaldehyde assimilation, showing the two variants for cleavage and the two variants for acceptor regeneration. Adapted from Anthony (1982).

Methylotrophic Bacteria Methanotrophs Methanotrophs are a subgroup of the methylotrophic bacteria, which have the ability to grow on methane as sole carbon and energy source. They are found in most environments in which methane and O2 meet, and have been isolated from a variety of environments including those with extremes of pH and temperature (Hanson and Hanson, 1996; Bodrossy et al., 1997; Bowman et al., 1997; Dedysh et al., 2000). They contain characteristic intracytoplasmic membrane systems (Hanson and Hanson, 1996), either stacks of membrane disks in the type I strains (g-Proteobacteria), rings of membranes at the periphery of the cell in the type II strains (Methylosinus and Methylocystis), or vesicular membranes in Methylocella (Dedysh et al., 2000). So far, all well-studied methanotrophs have been obligate methylotrophs, unable to grow on compounds with C–C bonds, but reports have been made of a facultative Mycobacterium strain capable of growth on methane (Reed and Dugan, 1987) and a mutant of a type I methanotroph has been described that is capable of growth on glucose (Zhao and Hanson, 1984). Some methanotrophs are capable of growth on methanol (Anthony, 1982; Hanson and Hanson,

1996). Some methanotrophs contain nitrogenase and are capable of growth with N2 as a nitrogen source, mainly Methylococcus and Methylosinus strains (Hanson and Hanson, 1996). Methanotrophs contain either the serine cycle or RuMP cycle, and so far no autotrophic methanotrophs have been identified (Table 1). Methanotrophs exist as symbionts in mussels, clams and Pogonophora, and although the 16S rRNA sequences class with type I methanotrophs, they have not yet been isolated in pure culture (Distel and Cavanaugh, 1994).

Non-Methane Utilizing Methylotrophs The bacteria capable of growth on methanol and other methylated compounds but not on methane are more diverse than those capable of growing on methane. So far, all of the Gram-positive and a-proteobacterial strains are facultative methylotrophs, whereas the b-proteobacterial and g-proteobacterial strains are either obligate methylotrophs or restricted facultative methylotrophs (capable of poor growth on a restricted range of multicarbon compounds; Anthony, 1982; Table 1). Most of these bacteria can grow on methanol and may grow on other methylated compounds, but a few strains are known that grow on methylated amines and in some cases also grow on other methylated compounds, but

CHAPTER 1.20

do not grow on methanol (Anthony, 1982). These include the Aminobacter (Urakami et al., 1992) and some of the strains that grow on halogenated methanes (Hancock et al., 1998). The nonmethane-utilizing methylotrophic bacteria do not generate intracytoplasmic membrane systems characteristic of the methanotrophs, with the exception of the photosynthetic membranes in the phototrophs. The phototrophs that grow on methanol use it as an electron donor for photosynthesis and in some cases as a carbon source (Quayle and Pfennig, 1975). A number of the Gram-negative methylotrophs contain nitrogenase and are capable of growth with N2 as a nitrogen source (Table 1). These bacteria are widely distributed in terrestrial, freshwater and marine habitats (Anthony, 1982). Bacteria capable of utilizing methylated amines are particularly prevalent in the marine environment where it is postulated that they may play a role in carbon cycling in the photic zone (Strand and Lidstrom, 1984). The pinkpigmented Methylobacterium strains are common epiphytes on plant leaves, and some evidence exists to suggest a mutualistic symbiosis (Holland and Polacco, 1994).

Genetics in Aerobic Methylotrophs Genetic Capabilities A variety of genetic capabilities are available in Gram-negative methylotrophic bacteria, mostly based on broad-host range vectors of the incompatibility (Inc)PI or IncQ groups, including both general cloning vectors and promoter probe vectors based on either lacZ or xylE as reporters (Holloway et al., 1987; Lidstrom and Sterling, 1990; Harms and van Spanning, 1991; De Vries et al., 1990; Barta and Hanson, 1993; Murrell et al., 2000). However, IncQ vectors are unstable in the Methylobacterium strains, and serve as suicide vectors (Biville et al., 1989). Targeted mutants can be generated from cloned genes by recombinational insertion in these strains using suicide vectors, most of them based on ColE1 replicons (Harms, 1996; Barta and Hanson, 1993; Chistoserdov and Lidstrom, 1994a; Murrell et al., 2000). The most common mode of transfer of these vectors into these methylotrophs is by conjugation using a helper plasmid, but electroporation protocols also have been reported for some of the nonmethanotrophs (Ueda et al., 1991; Kim and Wood, 1997; Gliesche et al., 1997; Toyama et al., 1998). So far, electroporation has not been successful for methanotrophs (Murrell et al., 2000). Random transposon mutagenesis has

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629

been reported for a number of Gram-negative methylotrophs (Whitta et al., 1985; Gliesche and Hirsch, 1992; Studer, 1999; Kang et al., 1999), but not for methanotrophs (Murrell et al., 2000). In the Gram-positive methylotrophs, cloning vectors have been developed for individual strains. The IncQ vectors and electroporation have been used for Brevibacterium methylicum (Nesvera et al., 1994), whereas in Bacillus methanolicus and Amycolatopsis methanolica, shuttle plasmids were developed using replicons from endogenous plasmids (Vrijbloed et al., 1995; Cue et al., 1997). In the latter two cases, genetic transformation systems were developed for vector transfer.

Genomics A number of genome sequencing projects are underway for methylotrophic bacteria, including Methylobacterium extorquens AM1, Methylococcus capsulatus Bath, Rhodopseudomonas palustris, Rhodobacter sphaeroides and Rhodobacter capsulatus.

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Strand, S. E., and M. E. Lidstrom. 1984. Characterization of a new marine methylotroph. FEMS Microbiol. Lett. 21:247–251. Studer, A., S. Vuilleumier, and T. Leisinger. 1999. Properties of the methylcobalamin:H4folate methyltransferase involved in chloromethane utilization by Methylobacterium sp. strain CM4. Eur. J. Biochem. 264:242–249. Suylen, G. M., and J. G. Kuenen. 1986. Chemostat enrichment and isolation of Hyphomicrobium EG: A dimethyl-sulphide oxidizing methylotroph and reevaluation of Thiobacillus MS1. Ant. v. Leeuwenhoek 52:281–293. Tanaka, Y., T. Yoshida, K. Watanabe, Y. Izumi, and T. Mitsunaga. 1997. Characterization, gene cloning and expression of isocitrate lyase involved in the assimilation of one-carbon compounds in Hyphomicrobium methylovorum GM2. Eur. J. Biochem. 249:820–825. Toyama, H., C. Anthony, and M. E. Lidstrom. 1998. Construction of insertion and deletion mxa mutants of Methylobacterium extorquens AM1 by electroporation. FEMS Microbiol. Lett. 166:1–7. Ueda, S., S. Matsumoto, S. Shimizu, and T. Yamane. 1991. Transformation of a methylotrophic bacterium, Methylobacterium extorquens, with a broad-host-range plasmid by electroporation. Ann. NY Acad. Sci. 646:99–105. Urakami, T., H. Araki, H. Oyanagi, K. I. Suzuki, and K. Komagata. 1992. Transfer of Pseudomonas aminovorans (den Dooren de Jong 1926) to Aminobacter, new genus as Aminobacter aminovorans, new subspecies and description of Aminobacter aganoensis, new species and Aminobacter niigataensis, new species. Int. J. Syst. Bacteriol. 42:84–92. van der Palen, C. J., D. J. Slotboom, L. Jongejan, W. N. Reijnders, N. Harms, J. A. Duine, and R. J. van Spanning. 1995. Mutational analysis of mau genes involved in methylamine metabolism in Paracoccus denitrificans. Eur. J. Biochem. 230:860–871. Vannelli, T., M. Messmer, A. Studer, S. Vuilleumier, and T. Leisinger. 1999. A corrinoid-dependent catabolic pathway for growth of a Methylobacterium strain with chloromethane. Proc. Natl. Acad. Sci. USA 96:4615–4620. Van Ophem, P. W., and J. A. Duine. 1990. Different types of formaldehyde-oxidizing dehydrogenases in Nocardia sp p. 239: Purification and characterization of an NADdependent aldehyde dehydrogenase. Arch. Biochem. Biophys. 282:248–253. Van Ophem, P. W., J. Van Beeumen, and J. A. Duine. 1992. NAD-linked, factor-dependent formaldehyde dehydrogenase or trimeric, zinc-containing, long-chain alcohol dehydrogenase from Amycolatopsis methanolica. Eur. J. Biochem. 206:511–518. Van Ophem, P. W., and J. A. Duine. 1994. NAD- and cosubstrate (GSH or factor)-dependent formaldehyde dehydrogenases from methylotrophic microorganisms act as a class III alcohol dehydrogenase. FEMS Microbiol. Lett. 116:87–93. Vorholt, J., L. Chistoserdova, M. E. Lidstrom, and R. K. Thauer. 1998. The NADP-dependent methylene tetrahydromethanopterin dehydrogenase in Methylobacterium extorquens AM1. J. Bacteriol. 180:5351–5356. Vorholt, J. A., L. Chistoserdova, S. M. Stolyar, R. K. Thauer, and M. E. Lidstrom. 1999. Distribution of tetrahydromethanopterin-dependent enzymes in methylotrophic bacteria and phylogeny of methenyl tetrahydromethanopterin cyclohydrolases. J. Bacteriol. 181:5750–5757.

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Vrijbloed, J. W., V. J. van Hylckama, N. M. van der Put, G. I. Hessels, and L. Dijkhuizen. 1995. Molecular cloning with a pMEA300-derived shuttle vector and characterization of the Amycolatopsis methanolica prephenate dehydratase gene. J. Bacteriol. 177:6666–6669. Walters, K. J., G. T. Gassner, S. J. Lippard, and G. Wagner. 1999. Structure of the soluble methane monooxygenase regulatory protein B. Proc. Natl. Acad. Sci. USA 96:7877–7882. Weaver, C. W., and M. E. Lidstrom. 1985. Methanol dissimilation in Xanthobacter H4-14: activities, induction and comparison to Pseudomonas AM1 and Paracoccus denitrificans. J. Gen. Microbiol. 131:2183–2197. Whitta, S., M. I. Sinclair, and B. W. Holloway. 1985. Transposon mutagenesis in Methylobacterium AM1 (Pseudomonas AM1). J. Gen. Microbiol. 131:1547–1549. Whittenbury, R., and H. Dalton. 1981. The methylotrophic bacteria. In: M. P. Starr, H. Stolp, H. G. Trüper, A. Balows, and H. G. Schlegel (Eds.) The Prokaryotes. Springer. New York, NY. 894–902. Yamada, Y., K. Hoshino, and T. Ishikawa. 1997. The phylogeny of acetic acid bacteria based on the partial sequences of 16S ribosomal RNA: the elevation of the subgenus Gluconoacetobacter to the generic level. Biosci. Biotechnol. Biochem. 61:1244–1251. Yanase, H., K. Ikeyama, R. Mitsui, S. Ra, K. Kita, Y. Sakai, and N. Kato. 1996. Cloning and sequence analysis of the gene encoding 3-hexulose-6-phosphate synthase from the methylotrophic bacterium, Methylomonas aminofa-

CHAPTER 1.20 ciens 77a, and its expression in Escherichia coli. FEMS Microbiol. Lett. 135:201–205. Yang, C. C., L. C. Packman, and N. S. Scrutton. 1995. The primary structure of Hyphomicrobium X dimethylamine dehydrogenase: Relationship to trimethylamine dehydrogenase and implications for substrate recognition. Eur. J. Biochem. 232:264–271. Yasueda, H., Y. Kawahara, and S. Sugimoto. 1999. Bacillus subtilis yckG and yckF encode two key enzymes of the ribulose monophosphate pathway used by methylotrophs, and yckH is required for their expression. J. Bacteriol. 181:7154–7160. Yoshida, T., K. Yamaguchi, T. Hagishita, T. Mitsunaga, A. Miyata, T. Tanabe, H. Toh, T. Ohshiro, M. Shimao, and Y. Izumi. 1994. Cloning and expression of the gene for hydroxypyruvate reductase (D-glycerate dehydrogenase) from an obligate methylotroph Hyphomicrobium methylovorum GM2. Eur. J. Biochem. 223:727–732. Zahn, J. A., and A. A. DiSpirito. 1996. Membrane-associated methane monooxygenase from Methylococcus capsulatus (Bath). J. Bacteriol. 178:1018–1029. Zatman, L. J. 1981. A search for patterns in methylotrophic pathways. In: H. Dalton (Ed.) Microbial Growth on C1 Compounds. Heyden. London, 42–54. Zhao, S. J., and R. S. Hanson. 1984. Variants of the obligate methanotroph isolate 761M capable of growth on glucose in the absence of methane. Appl. Environ. Microbiol. 48:807–812.

Prokaryotes (2006) 2:635–658 DOI: 10.1007/0-387-30742-7_21

CHAPTER 1.21 yrot a l imi s s iD

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

Introduction Dissimilatory Fe(III) reduction is the process in which microorganisms transfer electrons to external ferric iron [Fe(III)], reducing it to ferrous iron [Fe(II)] without assimilating the iron. A wide phylogenetic diversity of microorganisms, including archaea as well as bacteria, are capable of dissimilatory Fe(III) reduction. Most microorganisms that reduce Fe(III) also can transfer electrons to Mn(IV), reducing it to Mn(II). As detailed in the next section, dissimilatory Fe(III) and Mn(IV) reduction is one of the most geochemically significant events that naturally takes place in soils, aquatic sediments, and subsurface environments. Dissimilatory Fe(III) and Mn(IV) reduction has a major influence not only on the distribution of iron and manganese, but also on the fate of a variety of other trace metals and nutrients, and it plays an important role in degradation of organic matter. Furthermore, dissimilatory Fe(III)-reducing microorganisms show promise as useful agents for the bioremediation of sedimentary environments contaminated with organic and/or metal pollutants. Despite their obvious environmental significance, Fe(III) and Mn(IV)-reducing microorganisms are among the least studied of any of the microorganisms that carry out important redox reactions in the environment. The Fe(III)- and Mn(IV)-reducing microorganisms are also of intrinsically interesting because they have unique metabolic characteristics. Foremost is the ability of these microorganisms to transfer electrons to external, highly insoluble electron acceptors such as Fe(III) and Mn(IV) oxides, as well as extracellular organic compounds such as humic substances. Furthermore, microbiological and geological evidence suggests that dissimilatory Fe(III) reduction was one of the earliest forms of microbial respiration. Thus, insights into Fe(III) reduction mechanisms may aid in understanding the evolution of respiration in microorganisms.

Significance of Fe(III)- and Mn(IV)-Reducing Microorganisms Some claims for the significance of Fe(III)reducing microorganisms may be exaggerated, such as the assertion that “if it were not for the bacterium GS-15 [a Fe(III)-reducing microorganism] we would not have radio and television today” (Verschuur, 1993). However, it is also clear that Fe(III)-reducing microorganisms are of vitally important to the proper functioning of a variety of natural ecosystems and have practical applications. Detailed reviews of the literature covering many of these aspects of Fe(III) and Mn(IV) reduction are available (Lovley, 1987a; Lovley, 1991a; Lovley, 1993a; Nealson and Saffarini, 1994; Lovley, 1995a; Lovley et al. 1997c). Therefore only highlights of the significance of Fe(III)-reducing microorganisms, abstracted from these reviews, will be briefly summarized here.

Oxidation of Organic Matter in Anaerobic Environments Microbial oxidation of organic matter coupled to the reduction of Fe(III) and Mn(IV) is an important mechanism for organic matter oxidation in a variety of aquatic sediments, submerged soils, and in aquifers. Depending on the aquatic sediments or submerged soils considered, Fe(III) and/or Mn(IV) reduction have been estimated to oxidize anywhere from 10% to essentially all of the organic matter oxidation in the sediments (Lovley, 1991a; Canfield et al., 1993; Lovley, 1995b; Lovley et al., 1997c). An important factor that enhances the significance of Fe(III) and Mn(IV) reduction in aquatic sediments is bioturbation which leads to the reoxidation of Fe(II) and Mn(II) so that each molecule of iron and manganese can be used as an electron acceptor multiple times prior to permanent burial. In deep pristine aquifers, there are often extensive zones exist in which Fe(III) reduction is the predominant mechanism for organic matter oxidation

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(Chapelle and Lovley, 1992; Lovley and Chapelle, 1995c). The ability of Fe(III)-reducing microorganisms to outcompete sulfate-reducing and methanogenic microorganisms for electron donors during organic matter degradation is an important factor limiting the production of sulfides and methane in some submerged soils, aquatic sediments, and the subsurface (Lovley, 1991a; Lovley, 1995b). A model for the oxidation of organic matter in sedimentary environments in which Fe(III) reduction is the predominant terminal electronaccepting process has been suggested (Lovley et al., 1997c). This model is based upon the known physiological characteristics of Fe(III)and Mn(IV)-reducing microorganisms available in pure culture as well as on studies on the metabolism of organic matter metabolism by natural communities of microorganisms living in various sedimentary environments in which Fe(III) reduction is the terminal electron-accepting process (TEAP). In this model (Fig. 1), complex organic matter is hydrolyzed to simpler components by the action of hydrolytic enzymes from a variety of microorganisms. Fermentative microorganisms are the principal consumers of fermentable compounds such as sugars and amino acids and these compounds are converted primarily to fermentation acids and, possibly to hydrogen. Acetate is by far the most important fermentation acid produced (Lovley and Phillips, 1989a). Acetate also may be produced as the result of incomplete oxidation of some sugars by some Fe(III)-reducing microorganisms (Coates et al., 1999a). Other Fe(III)-reducing microorganisms oxidize the acetate and other intermediary products. Some Fe(III)-reducing microorganisms also can oxidize aromatic compounds and long-chain fatty acids. Thus, through the activity of diverse microorganisms, complex organic matter can be oxidized to carbon dioxide with Fe(III) serving as the sole electron acceptor. A similar model probably is probably appropriate for organic matter oxidation in sediments in which Mn(IV) reduction is the TEAP. This

Aromatic Compounds hilia

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She Hydrolysis of Complex Organic Matter

Fermentable Substrates

Fermentative Microorganisms

Long Chain Fatty Acids

model emphasizes that acetate is likely to be the major electron donor for Fe(III) or Mn(IV) reduction in environments in which naturally occurring, complex organic matter is the major substrate for microbial metabolism. However, when otherwise organic-poor environments, such as sandy aquifers, are contaminated with a specific class of organic compounds, such as aromatics, then these contaminants may be the most important direct electron donors for Fe(III) or Mn(IV) reduction. Influence on Metal and Nutrient Geochemistry and Water Quality The reduction of Fe(III) to Fe(II) is one of the most important geochemical changes as anaerobic conditions develop in submerged soils and aquatic sediments (Ponnamperuma, 1972). The Fe(II) produced as the result of Fe(III) reduction is the primary reduced species responsible for the negative redox potential in many anaerobic freshwater environments. The reduction of Fe(III) oxides and of the structural Fe(III) in clays typically results in a change in soil color from the redyellow of Fe(III) forms to the green-gray of Fe(II) minerals (Lovley, 1995c). The oxides of Fe(III) and Mn(IV) oxides bind trace metals, phosphate, and sulfate, and Fe(III) and Mn(IV) reduction is associated with the release of these compounds into solution (Lovley, 1995a). Also, typically the pH, ionic strength of the pore water, and the concentration of a variety of cations are increased (Ponnamperuma, 1972; 1984). All of these changes influence water quality in aquifers and can affect the growth of plants in soils. The solubility of Fe(II) and Mn(II) is greater than that of Fe(III) and Mn(IV) and thus Fe(III) and Mn(IV) reduction result in an increase in dissolved iron and manganese in pore waters. Undesirably high concentrations of iron and manganese may be toxic to plants (Lovley, 1995b) and are particularly significant in groundwaters sources of drinking water, being one of the most prevalent groundwater quality problems (Anderson and Lovley, 1997).

Ge

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rm ata ller edu Ge cen o Acetate Ge bac s ov te Minor ibr r, G i a e o fermentation Fe th rri rix acids ac ier othrix ter, Ge eobac H2 G Shewanella ans itatis ont ferm s patm x i a r th cn Geo lfurom u Dex

Fe(II) CO2

Fig. 1. Proposed pathways for organic matter degradation in mesophilic environments in which Fe(III) reduction is the predominant terminal electron-accepting process.

CHAPTER 1.21

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

Most of the Fe(II) and Mn(II) produced from microbial Fe(III) and Mn(IV) reduction is found in solid phases, often in the form of Fe(II) and Mn(II) minerals of geochemical significance (Lovley, 1995c). The most intensively studied mineral that is formed during microbial Fe(III) reduction is the magnetic mineral magnetite (Fe3O4) (Lovley et al., 1987c; Lovley, 1990a; Lovley, 1991a). The magnetite produced during microbial Fe(III) reduction can be an important geological signature of this activity. For example, large quantities of magnetite at depths up to 6.7 km below the Earth’s surface provided some of the first evidence for a deep, hot biosphere (Gold, 1992). The massive magnetite accumulations that comprise the Precambrian Banded Iron Formations provide evidence for the possible activity of Fe(III)-reducing microorganisms on early Earth. Formation of magnetite as the result of microbial Fe(III) reduction may contribute to the magnetic remanence of soils and sediments. The magnetic anomalies that aid in the localization of subsurface hydrocarbon deposits may result from the activity of hydrocarbon-degrading Fe(III) reducers. Formation of other Fe(II) and Mn(II) minerals such as siderite (FeCO3) and rhodochrosite (MnCO3) also may provide geological signatures of microbial Fe(III) and Mn(IV) reduction. As detailed below, many Fe(III)- and Mn(IV)reducing microorganisms can use other metals and metalloids as electron acceptors. Microbial reduction of the soluble oxidized form of uranium, U(VI), to insoluble U(IV) may be an important mechanism for the formation of uranium deposits and the reductive sequestration of uranium in marine sediments, the process which prevents dissolved uranium from building up in marine waters (Lovley et al., 1991a; Lovley and Philips, 1992). Reduction of other metals such as vanadium, molybdenum, copper, gold, and silver, as well as metalloids such as selenium and arsenic, can affect the solubility and fate of these compounds in a variety of sedimentary environments and may contribute to ore formations (Lovley, 1993a; Oremland, 1994a; Newman et al., 1998; Kashefi and Lovley, 1999). Bioremediation of Organic and Metal Contaminants Iron [Fe(III)]-reducing microorganisms have been shown to play a major role in removing organic contaminants from polluted aquifers. For example, Fe(III)-reducing microorganisms naturally remove aromatic hydrocarbons from petroleum-contaminated aquifers (Lovley et al., 1989b; Lovley, 1995c; Lovley, 1997a; Anderson et al., 1998) and this process can be artificially enhanced with compounds that make Fe(III) more available for microbial reduction (Lovley et al., 1994a; Lovley, 1997a). The

637

Fe(II)-minerals formed as the result of microbial Fe(III) reduction can be important reductants for the reduction of nitroaromatic contaminants (Heijman et al., 1993; Hofstetter et al., 1999). Minerals containing Fe(II) also may serve to reductively dechlorinate some chlorinated contaminants (Fredrickson and Gorby, 1996). The ability of Fe(III)-reducing microorganisms to substitute other metals and metalloids in their respiration may be exploited for remediation of metal contamination (Lovley, 1995a; Lovley, 1995b; Fredrickson and Gorby, 1996; Lovley and Coates, 1997b). Reduction of soluble U(VI) to insoluble U(IV) can effectively precipitate uranium from contaminated groundwaters and surface waters. Microbial uranium reduction can be coupled with a simple soil-washing procedure to concentrate uranium from contaminated soils. Iron [Fe(III)]-reducing microorganisms can precipitate technetium from contaminated waters by reducing soluble Tc(VII) to insoluble Tc(IV). Soluble radioactive Co(III) complexed to EDTA can be reduced to Co(II) which is less likely to be associated with the EDTA found in contaminated groundwaters and more likely to adsorb to aquifer solids. Some Fe(III) reducers convert soluble, toxic Cr(VI) to less soluble less toxic Cr(III). Reduction of soluble selenate to elemental selenium can effectively precipitate selenium in sediments or remove selenate from contaminated waters in bioreactors. A Possible Early Form of Microbial Respiration Iron [Fe(III)] reduction may have been one of the earliest forms of microbial respiration (Vargas et al., 1998). Biological evidence for this hypothesis is the finding from 16S rRNA phylogenies that all of microorganisms that are the most closely related to the last common ancestor of extant microorganisms are Fe(III)reducing microorganisms. All of the deeply branching bacteria and archaea that have been examined can oxidize hydrogen with the reduction of Fe(III). Several that have been examined in more detail can conserve energy to support growth from this metabolism. Of most interest in this regard is Thermotoga maritima, which was previously considered to be a fermentative organism because it could not conserve energy to support growth from the reduction of other commonly considered electron acceptors. However, T. maritima it does grow via Fe(III) respiration. This result and the apparent conservation of the ability to reduce Fe(III) in all these deeply branching organisms suggests that the last common ancestor was a hydrogen-oxidizing, Fe(III)reducing microorganism. The concept that Fe(III) reduction is an early form of respiration agrees with geological scenarios that suggest the presence of large quanti-

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D. Lovley

ties of Fe(III) on prebiotic Earth (Cairns-Smith et al., 1992; de Duve, 1995) and elevated hydrogen levels (Walker, 1980)—conditions that would be conducive to the evolution of a hydrogen-oxidizing, Fe(III)-reducing microorganism. The large accumulations of magnetite in the Precambrian iron formations (discussed above) indicate that the accumulation of Fe(III) on prebiotic Earth was biologically reduced early in the evolution of life on Earth. This and other geochemical considerations suggest that Fe(III) reduction was the first globally significant mechanism for organic matter oxidation (Walker, 1987; Lovley, 1991a).

Fe(III)- and Mn(IV)-Reducing Microorganisms Available in Pure Culture Dissimilatory Fe(III)- and Mn(IV)-reducing microorganisms can be separated into two major groups, those that support growth by conserving energy from electron transfer to Fe(III) and Mn(IV) and those that do not. Early investigations on Fe(III) and Mn(IV) reduction in pure culture were conducted exclusively with organisms that are not considered to be conservers of energy from Fe(III) or Mn(IV) reduction (Lovley, 1987a). However, within the last decade, a diversity of microorganisms has been described in which Fe(III) and Mn(IV) reduction are linked to respiratory systems capable of ATP generation. It is these Fe(III)- and Mn(IV)respiring microorganisms (abbreviated here as FMR) that are likely to be responsible for most of the Fe(III) and Mn(IV) reduction in many sedimentary environments (Lovley, 1991a). A brief description of the known metabolic and phylogenetic diversity of dissimilatory Fe(III)and Mn(IV)-reducing microorganisms follows. Fermentative Fe(III)- and Mn(IV)-Reducing Microorganisms Many microorganisms which grow via fermentative metabolism can use Fe(III) or Mn(IV) as a minor electron acceptor during fermentation (Table 1). Growth is possible in the absence of Fe(III) or Mn(IV). In this form of Fe(III) and Mn(IV) reduction, most of the electron equivalents in the fermentable substrates are recovered in organic fermentation products and hydrogen. Typically, less than 5% of the reducing equivalents are transferred to Fe(III) or Mn(IV) (Lovley, 1987a; Lovley and Phillips, 1988b). However, significant amounts of Fe(II) and Mn(II) can accumulate in cultures of these fermentative organisms when Fe(III) or Mn(IV) is provided as a potential electron sink. Although thermodynamic calculations have demonstrated that fermentation with Fe(III) reduction [electron transfer to Fe(III)] is more energetically favorable than fermentation with-

CHAPTER 1.21

out Fe(III) reduction (Lovley and Phillips, 1989a), it has not been demonstrated that the minor transfer of electron equivalents to Fe(III) or Mn(IV) during fermentation causes any increase in cell yield. In contrast to these fermentative microorganisms, several microorganisms can partially or completely oxidize fermentable sugars and amino acids with the reduction of Fe(III) and conserve energy from this metabolism, as discussed below. Sulfate-Reducing Microorganisms Many respiratory microorganisms that grow anaerobically with sulfate serving as the electron acceptor also have the ability to enzymatically reduce iron [Fe(III); Table 1]. Electron donors that support Fe(III) reduction are the same ones that support sulfate reduction by sulfate-reducing microorganisms. However, none of these sulfate reducers have been shown to grow with Fe(III) serving as the sole electron acceptor (Lovley et al., 1993b). This is true despite the fact that sulfate reducers have a higher affinity for hydrogen, and possibly for other electron donors, than for sulfate when Fe(III) serves as the electron acceptor (Coleman et al., 1993; Lovley et al., 1993c). The advantage to sulfate reducers in reducing Fe(III), if there is one, has not been thoroughly investigated. Because it has been found that the intermediate electron carrier, cytochrome c3, can function as an Fe(III) reductase (Lovley et al., 1993), intermediate electron carriers involved in sulfate reduction may inadvertently reduce Fe(III) because it has been found that the intermediate electron carrier, cytochrome c3 can function as an Fe(III) reductase (Lovley et al., 1993b). Alternatively, Fe(III) reduction by sulfate reducers may be a strategy to hasten Fe(III) depletion and enhance conditions for sulfate reduction. Furthermore, the possibility that sulfate-reducing microorganisms may be able to generate ATP as the result of Fe(III) reduction, even if they can not grow with Fe(III) as the sole electron acceptor, has not been ruled out (Lovley et al., 1993c). In contrast to the sulfate-reducing microorganisms discussed above, which could not be grown with Fe(III) as the sole electron acceptor, it has been suggested (Tebo and Obraztsova, 1998) that the sulfate-reducing microorganism “Desulfotomaculum reducens” could also conserve energy to support growth by reducing Fe(III), Mn(IV), U(VI), and Cr(VI) (Tebo and Obraztsova, 1998). However, the data supporting the claim that energy is gained from electron transport to metals is curious. For example, when the culture was grown on 400 mM U(VI), the cell yield was greater than when the culture reduced 8 mmol Fe(III). This occurs despite the fact that the number of electrons transferred to Fe(III)

CHAPTER 1.21

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

639

Table 1. Organisms known to reduce Fe(III) but not known to conserve energy from Fe (III) reduction. Electron donor

Form of Fe(III) reduceda

Reference

Bacillus pumilus Bacillus sp. Bacillus subtilis Bacteroides hypermegas Clostridium butyricum Clostridium polymyxa Clostridium saccarobutyricum Clostridium sporogenes Escherichia coli Escherichia coli Fusarium oxysporum Fusarium oxysporum Fusarium solani Paracolobactrum sp. Pseudomonas aeruginosa Pseudomonas denitrificans Pseudomonas liquefaciens Pseudomonas (several species) Rhodobacter capsulatus Serratia marcescans Sulfolobus acidocaldarius Thiobacillus thiooxidans

Glucose Glucose-asparagine Glucose Glucose Glucose-asparagine Sucrose Glucose-asparagine Sucrose Sucrose Glucose Glucose Sucrose Glucose-asparagine Glucose Glucose Glucose-tryptone Glucose Sucrose Glucose Glucose, peptone Glucose, peptone Glucose-asparagine Glucose Glucose Glucose Glucose Glucose-asparagine Glucose Sucrose Glucose-asparagine Malate Glucose-asparagine Elemental sulfur Elemental sulfur

Hematite Hematite PCIO Hematite Hematite PCIO Hematite Ferro-manganese ore Ferro-manganese ore PCIO Hematite PCIO Hematite Limonite, goethite, hematite Hematite Fe(III)-Cl3 Hematite Ferro-manganese ore Hematite PCIO PCIO Hematite Ferric ammonium citrate Hematite Hematite PCIO Hematite Fe(III)-Cl3 Ferro-manganese ore Hematite Fe(III)-NTA Hematite Fe(III)-Cl3 Fe(III)-Cl3

Vibrio sp. Vibrio sp. Wolinella succinogenes

Glucose Malate, pyruvate Formate

Fe(III)-Cl3 Fe(III)-Cl3 Fe(III)-Cit

Ottow and von Klopotek, 1969 Ottow, 1970 Bromfield, 1954 Ottow and von Klopotek, 1969 Ottow, 1970 Bromfield, 1954 Ottow, 1970 Troshanov, 1968 Troshanov, 1968 Roberts, 1947 Hammann and Ottow, 1974 Bromfield, 1954 Ottow, 1970 De Castro and Ehrlich, 1970 Ottow and Glathe, 1971 Jones et al., 1984a Hammann and Ottow, 1974 Troshanov, 1968 Hammann and Ottow, 1974 Starkey and Halvorson, 1927 Starkey and Halvorson, 1927 Ottow, 1970 Gunner and Alexander, 1964 Ottow and von Klopotek, 1969 Ottow and von Klopotek, 1969 Bromfield, 1954 Ottow, 1970 Jones et al., 1984a Troshanov, 1968 Ottow and Glathe, 1971 Dobbin et al., 1996 Ottow, 1970 Brock and Gustafson, 1976 Brock and Gustafson, 1976 Kino and Usami, 1982 Jones et al., 1983 Jones et al., 1984b Lovley et al., 1998

Fe(III)-NTA Fe(III)-NTA Fe(III)-NTA Fe(III)-NTA

Lovley et al., 1993 Lovley et al., 1993 Lovley et al., 1993 Lovley et al., 1993

Desulfovibrio desulfuricans

Acetate H2 Propionate Butyrate, caproate, octanoate Lactate

Fe(III)-Cl3

Desulfovibrio baculatus Desulfovibrio sulfodismutans Desulfovibrio vulgaris Desulfotomaculum nigrificans

Lactate Lactate Lactate Lactate

Fe(III)-NTA Fe(III)-NTA Fe(III)-NTA Fe(III)-Cl3

Jones et al., 1984a Coleman et al., 1993 Lovley et al., 1993 Lovley et al., 1993 Lovley et al., 1993 Jones et al., 1984a

Archaea Archaeoglobus fulgidus Methanococcus thermolithotrophicus Methanopyrus kandleri Pyrococcus furiosus Pyrodictium abyssi

H2 H2 H2 H2 H2

Fe(III)-Cit Fe(III)-NTA Fe(III)-Cit Fe(III)-Cit Fe(III)-Cit

Vargas et al., 1998 Vargas et al., 1998 Vargas et al., 1998 Vargas et al., 1998 Vargas et al., 1998

Organism Fermentative bacteria Actinomucor repens Aerobacter aerogenes Aerobacter sp. Alternaria tenuis Bacillus cereus Bacillus circulans Bacillus mesentericus Bacillus polymyxa

Sulfate-reducing bacteria Desulfobacter postgatei Desulfobacterium autotrophicum Desulfobulbus propionicus Desulfovibrio baarsii

a

Fe(III) forms: Poorly crystalline Iron Oxide (PCIO), Ferric citrate [Fe(III)-Cit], Ferric nitriloacetic acid [Fe(III)-NTA], Ferric pyrophosphate [Fe(III)-P], Fe(III) chloride [Fe(III)-Cl3].

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CHAPTER 1.21

was ten-fold higher than the electron transfer to U(VI) and that Fe(III) reduction is energetically more favorable than U(VI) reduction. Cell yields with metals as the electron acceptor were comparable to those during sulfate reduction even though electron transfer to sulfate was at least 250-fold, and in some instances 2500-fold, greater than electron transfer to the metals. These results suggest that the presence of the metals had some additional influence on growth other than just serving as an electron acceptor. Several sulfate-reducing microorganisms can oxidize S∞ to sulfate, with Mn(IV) serving as the electron acceptor, but were not found to conserve energy to support growth from this reaction (Lovley and Phillips, 1994a). Enrichment cultures that are established at circumneutral pH with S∞ as the electron donor and Mn(IV) or Fe(III) as the electron acceptor typically yield microorganisms which that disproportionate S∞ to sulfate and sulfide (Thamdrup et al., 1993). The Fe(III) or Mn(IV) serve to abiotically reoxidize the sulfide produced.

Microorganisms that Conserve Energy to Support Growth from Fe(III) and Mn(IV) Reduction The Fe(III)- and Mn(IV)-respiring microorganisms (FMR) which are known to conserve energy to support growth from Fe(III) and Mn(IV) reduction (Table 2) are phylogenetically (Fig. 2)

and morphologically (Fig. 3) diverse. Most of the FMR grow by oxidizing organic compounds or hydrogen with the reduction of Fe(III) or Mn(IV), but S∞ oxidation coupled to Fe(III) reduction also can provide energy to support growth of microorganisms growing at low pH. The various types of FMR are briefly described below. Geobacteraceae Most of the known FMR, available in pure culture, that can oxidize organic compounds completely to carbon dioxide with Fe(III) or Mn(IV) serving as the sole electron acceptor are in the family Geobacteraceae in the delta d-Proteobacteria (Fig. 2; Table 2). The family Geobacteraceae is comprised of the genera Geobacter, Desulfuromonas, Desulfuromusa and Pelobacter. With the exception of the Pelobacter species, all of the Geobacteraceae genera contain microorganisms that oxidize acetate to carbon dioxide. This metabolism is significant because, as discussed above, acetate is probably the primary electron donor for Fe(III) reduction in most sedimentary environments. Many of these Geobacteraceae also can use hydrogen as an electron donor for Fe(III) reduction. Various species in the Geobacteraceae oxidize a variety of other organic acids, including in some instances long-chain fatty acids (Table 2). Several species of Geobacter have the ability to anaerobically oxidize aromatic compounds, including the hydrocarbon toluene.

0.1 substitutions/site 100

Desulfuromonas chloroethenica Desulfuromonas aetexigens Pelobacter renetianus Pelobacter carbinolicus Desulfuromonas polmitatis Desulfuromonas succinoxidans 100 Desulfuromonas bakii 70 Desulfuromonas kyisingii 100 Desulfuromonas acetoxidans Pelobacter proplonicus 100 Geobacter chapellel Geobacter sulfurreducens 100 60 Geobacter hydrogenopiholus 99 100 Geobacter metallireducens 95 Geobacter grbicium Sulfurospirillum barnesii Thiobacillus ferrooxidans 65 Ferribacterium limneticum 94 97 87 Ferrimanas balearica Aeromonas hydrophila 100 Shewanella saccharophilia 100 Shewanella putrefaciens Shewanella alga Geobacter fementans 62 30

Deferribacter themophilus Geovibria femreucens Bacillus infenus Themnoterrahacterium ferrireducens Themus sp.SA-01 Thenmotoga marthma 100

97 100 100

100

Pyrobaculum aerophilum Pyrobaculum islandicum

Fig. 2. Phylogenetic tree, based on 16S rDNA sequences, of microorganisms known to conserve energy to support growth from Fe(III) reduction. The tree was inferred using the Kimura two-parameter model in TREECON for Windows (Van der Peer and De Wachter, 1994). Bootstrap values at nodes were calculated from one hundred replicates.

Freshwater ditch

Deep subsurface

“Geobacter arculus”

“Geobacter chapellei” (strain 172) “Geobacter grbicium” (strain TACP-2)

Aquatic sediments

Freshwater ditch

Mine-impacted lake sediments Marine sediments

Marine and freshwater muds Freshwater anoxic muds Marine sediments

“Geobacter akaganeitreducens”

Desulfuromusa kysingii Desulfuromusa succinoxidans Ferribacterium limneticum Ferrimonas Balearica

Desulfuromusa bakii

Marine sediments

ND

Lac

Ac, Buty, EtOH, For, Prop, Pyr

Complete

Ac, EtOH, For, Fum, ND H2, Mal, Prop, PrOH, Pyr, Succ Ac, BtOH, Buty, Bzo, ND EtOH, For, Fum, H2, Lac, Mal, Prop, PrOH, Pyr, Succ Ac, EtOH For, Lac Complete

Complete

Complete

Ac Ac

Complete

Complete

Complete

ND

Ac

Ac

Ac, Fum, Lac, Lau, Pal, Ste, Succ

Marine sediments

Freshwater sediments

Ac, BtOH, EtOH Prop, Pyr Ac, Pyr

Anoxic muds

Desulfuromonas acetexigens Desulfuromonas acetoxidans Desulfuromonas chloroethenica Desulfuromonas palmitatis Complete

For, Lac Incomplete NDf Ac, CAA, H2, Lac, Mal, Pept, Pyr, Succ, Try, Valr, YE Ac Complete

Deep subsurface North Sea oil field

Complete

Bacillus infernus Deferribacter thermophilus

Glyc, Lac, Succ

Freshwater and sewage

Source

Oxidation with Fe(III)b

Aeromonas hydrophila

Organism

Electron donors oxidized with Fe(III)a

Table 2. Organisms known to conserve energy to support growth from Fe(III) reduction.

PCIO, Fe-NTA, PCIO, Fe(III)-Cit

PCIO

PCIO, Fe(III)-P PCIO, Fe(III)-Cit PCIO, Akaganeite

Fe(III)-Cit, Fe(III)-NTA Fe(III)-NTA

PCIO, Fe(III)-Cit, Fe(III)-NTA, Fe(III)-P Fe(III)-NTA

Fe(III)-Cit, Fe(III)-NTA Fe(III)-NTA

PCIO

PCIO, Fe(III)-Cit Fe(III)-Cl3 PCIO, Fe(III)-Cit

Fe forms reducedc

AQDS

30

25

30

Mn(IV), S0, Fum, Mal

Mn(IV), AQDS, Fum

30

Mn(IV), So, Fum, Mal

Rod

Rod

Rod

Rod

Rod

37 Mn(IV), Nitrate

Rod

Rod

Rod

Rod

Rod

Rod

Rod

30

30

25

40

21–31

30

Rod

Rod Rod

60 60 30

Rod

Morphology

37

Growth Temp (∞C)

25

Nitrate, Fum

S∞, Mal, Fum, DMSO, nitrate S∞, Mal, Fum

S∞, Mal, Fum

Mn(IV), AQDS, S0, Fum

PCE, TCE, Fum, S2-

Mn(VI), S∞, polysulfides, Fum, Mal Mn(IV), Glut, Mal, Fum

U(VI), Co(III), selenate, nitrate, Fum, O2 Mn(IV), nitrate, TMAO Mn(VI), nitrate

Other electron acceptorsd

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

Coates et al., 1996

Coates et al., 1996

Straub et al., 1998

Straub et al., 1998

Rossello-Mora et al., 1995

Cummings et al., 1999

Liesack and Finster, 1994 Lonergan et al., 1996 Lonergan et al., 1996

Lonergan et al., 1996

Coates et al., 1995

Krumholz, 1997

Roden and Lovley, 1993

Coates et al., 1995

Knight and Blakemore, 1998 Boone et al., 1995 Greene et al., 1997

Referencee

CHAPTER 1.21 641

Contaminated aquifer

Contaminated ditch

Marine sediments

Geothrix Fermentans

Geovibrio Ferrireducens

Pelobacter Carbinolicus Pelobacter Propionicus Pelobacter Venetianus “Pseudomonas sp.”

Geothermal water

Marine hydrothermal waters Estuarine Sediment

Pyrobaculum Islandicum

Pyrobaculum Aerophilum Shewanella alga

Freshwater sediments Swampy soil

Contaminated ditch

Incomplete Incomplete ND

ND Incomplete

Lac EtOH, For, H2 H2 H2, Pept, YE

H2, Pept, YE H2, Lac

Incomplete

Complete

Ac, CAA, Fum, H2, Lac, Pro, Prop, Pyr, Succ, YE EtOH, H2

Mn(VI), U(VI)*, S2O32-, AQDS, TMAO, Fum, O2

Mn(IV)*, U(VI)*, Co(III)*, Tc(VII)*, Cr(VI)*, Au(III)*, Cyst, Glut, S0, SO32-, S2O32Nitrate, nitrite, O2

S∞ S∞ Nitrate, O2

S∞

Co(III), S∞

Mn(IV), AQDS, S∞

Tc(VII)*, Co(III), AQDS, S∞, Fum, Mal

Mn(IV), Tc(VII)*, U(VI), AQDS, humics, Nitrate

AQDS, Fum

Mn(IV), AQDS S∞, nitrate, Fum,

Other electron acceptorsd

30

100

100

30 30

30

35

30

35

30

30

30

Growth Temp (∞C)

Rod

Rod

Rod

Rod Rod Rod

Rod

Vibrio

Rod

Rod

Rod

Rod

Rod

Morphology

Cacavvo et al., 1992

Kashefi et al., 1999

Lonergan et al., 1996 Lonergan et al., 1996 Balashova and Zavarzin, 1980 Kashefi et al., 1999

Lovley et al., 1995

Cacavvo et al., 1996

Coates et al., 1999

Cacavvo et al., 1994

Lovley and Philips, 1988; Lovley et al., 1993

Coates et al., 1996

Coates et al., 1998

Referencee

D. Lovley

PCIO, Fe(III)-cit PCIO, Fe(III)-cit

PCIO, Fe(III)-cit

Fe(III)-NTA Fe(III)-NTA PCIO

PCIO, Fe(III)-Cit, Fe(III)-P PCIO, Fe(III)-Cit, Fe-NTA PCIO, Fe(III)-citrate, Fe(III)-P Fe(III)-NTA

Complete Complete

PCIO, Fe(III)-Cit

PCIO, Fe(III)-Cit

Complete Complete

PCIO, Fe(III)-Cit

Fe forms reducedc

Complete

Oxidation with Fe(III)b

Ac, Lac

Ac, Buty, Bzo, EtOH, For, H2, Prop, Pyr, Suc Ac, Bz, BzOH, BtOH, Buty, Bzo, BzOH, p-CR, EtOH, p-HBz, p-HBzo, p-HBzOH, IsoB, IsoV, Ph, Prop, PrOH, Pyr, Tol, Valr Ac, For, H2

Contaminated aquifer

Aquatic sediments

Ac, EtOH, For, H2, Lac

Electron donors oxidized with Fe(III)a

Contaminated wetland

Source

Geobacter sulfurreducens

“Geobacter humireducens” (strain JW3) “Geobacter hydrogenophilus” (strain H2) Geobacter metallireducens

Organism

Table 2. Continued

642 CHAPTER 1.21

Geothermally heated sea floor Hot springs, Yellowstone Deep gold-mine Groundwater

Mine drainage

Thermotoga Maritima

Thiobacillus Ferrooxidans

Incomplete

For, EtOH, H2, Lac, Mal

ND Incomplete ND ND

H2, Glyc Lac S0

Incomplete

H2

For, H2, Lac

For, Glyc, H2, Lac, Pyr, Incomplete Suc, YE

Incomplete

For, H2, Lac Pyr

Oxidation with Fe(III)b

Fe2(SO4)3

PCIO, Fe(III)-Cit Fe(III)-Cit, Fe(III)-NTA

Fe(III)-Cit

PCIO, Fe(III)-cit, Fe-NTA, Fe(III)-P, Fe(III)-EDTA PCIO, Fe(III)-cit

PCIO Fe(III)-cit

PCIO, Fe(III)-cit

Fe forms reducedc

30

Rod

65

Mn(VI), Co(III)*, Cr(VI)*, U(VI)*, S0, AQDS, nitrate, O2 S∞, O2,

Rod

65

AQDS, S2O32-, Fum

Rod

Rod

Vibrio

Rod

Rod

Rod

Morphology

80

30

30

30

30

Growth Temp (∞C)

Mn(IV), selenate, arsenate, S2O32-, S∞, nitrite, nitrate, Fum. TMAO, O2 S0

Mn(IV), U(VI) *, S∞, AQDS, S2O32-, nitrate, Mal, Fum, O2

Mn(VI), U(VI), S∞, S2O32-, AQDS, nitrate, Fum, O2 Mn(IV), SO32-, Nitrate, Fum, TmaO

Other electron acceptorsd

Das et al., 1992 Pronk et al., 1992

Kieft et al., 1999

Slobodkin et al., 1997

Vargas et al., 1998

Laverman et al., 1995

Coates et al., 1998

D. Boone personal communication

Myers and Nealson, 1988; Lovley et al., 1989

Referencee

a Abbreviations for electron donors and acceptors: Acetate(Ac), Anthraquinone-2,6-disulfonic acid (AQDS), Alanine(Ala), Aspartate (Asp), Benzaldehyde (Bz), Benzoate(Bzo), Benzylalcohol (BzOH), 1,2 butanediol (1,2 Bu), Butanol (BtOH), Butyrate (Buty), Casamino acids (CAA), Casein (Cas), Cystine (Cyst), Dimethylsulfoxide (DMSO), Ethanol (EtOH), Formate (For), Fumarate (Fum), Gelatin (GE), Glucose (Glu), Glutamate (Glu), Glutathione, oxidized (Glut), Glycerol (Glyc), p-hydroxybenzoate (p-HB), p-hydroxybenzaldehyde (p-HBz), phydroxybenzylalcohol (p-HBzOH), p-cresol (p-Cr), Hydrogen (H2), Inositol (Ino), Isobutyrate (IsoB), Isovalerate (IsoV), Lactate (Lac), Laurate (Lau), Malate (Mal), Maleate (Mle), Nitriloacetic acid (NTA), Oxaloacetate (OAA), Palmitate (Pal), Peptone (Pept), Phenol (Ph), Proline (Pro), Propanol (PrOH), Propionate (Prop), Pyruvate (Pyr), Ribose (Rib), Stearate (Ste), Succinate (Succ), Sucrose (Suc), Tetrachloroethylene (PCE), Trichloroethylene TCE), Toluene (Tol), Trimethylene oxide (To), Trimethylamine oxide (TMAO), Tryptone (Try), Valerate (Valr), Yeast extract (YE), Xylose (Xyl). b Complete oxidation of multicarbon compounds to CO2, or incomplete, typically to acetate. c Fe(III) forms: Poorly crystalline iron oxide (PCIO), ferric citrate [Fe(III)-cit]; ferric nitriloacetic acid, [Fe(III)-NTA]; ferric pyrophosphate, [Fe(III)-P]; Fe(III) chloride, [Fe(III)-Cl3]; Fe(III) ethylenediamine-tetraacetic acid. d * Organism has the ability to reduce the metal but not determined whether energy to support growth is conserved from reduction of this metal. e Reference in which the capacity to grow via Fe(III) reduction is described. f ND = Not determined. Superscript for electron acceptors if growth occurs. Coates, J.D., Council, T., Ellis, D.J., 1999. Carbohydrate oxidation coupled to Fe(III) reduction—a novel form of anaerobic metabolism. Anaerobe 4 277–282. Coates J. D., Bhupathivaju V., Achenbach L.A., McInerney M.J., Geobacter hydrogenophilus, Geobacter chapellii, Geobacter grbicium—three new strictly anaerobic dissimilatory Fe(III)reducers IJSB submitted. Coates Ellis Gaw Lovely. Geothrix fermentans gen. nov. sp. nov. a novel Fe(III) reducing bacterium from a hydrocarbon contaminated aquifer IJSB in press. Patrick J. A., Achenbach L.A., and Coates J. D. 1999. Geobacter humireducens—Eight new humic-reducing bacteria from a diversity of environments. IJSB submitted.

Thermoterrabact. Ferrireducens Thermus strain SA-01

Freshwater marsh

Aquatic Sediments

Aquatic sediments and other diverse environments Subsurface

Source

Sulfurospirillum Barnesii

“Shewanella putrefaciens CN32” Shewanella Saccharophilia

Shewanella Putrefaciens

Organism

Electron donors oxidized with Fe(III)a

CHAPTER 1.21 Dissimilatory Fe(III)- and Mn(IV)-Reducing Prokaryotes 643

644

D. Lovley

CHAPTER 1.21

Geobacter metallireducens

Geobacter sulfurreducens

Desulfuromonas palmitatis

Geothrix fermentans

Geovibrio ferrireducens

Ferribacterium limneticum

Shewanella alga

Bacillus infemus

Pyrobaculum iskindicum

Fig. 3. Phase contrast micrographs of various organisms that conserve energy to support growth from Fe(III) reduction. Bar equals 5 mm, all micrographs at equivalent magnification.

Geobacteraceae are the Fe(III) reducers most commonly recovered from a variety of sedimentary environments when the culture media contains acetate as the electron donor and Fe(III) oxide or the humic acid analog, anthraquinone2,6-disulfonate (AQDS) as the electron acceptor

(Coates et al., 1996; Coates et al., 1998). Furthermore, analysis of 16S rDNA sequences in sandy aquifer sediments in which Fe(III) reduction was the predominant terminal electron accepting process indicated that Geobacter species were a major component of the microbial community

CHAPTER 1.21

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

645

(Rooney-Varga et al., 1999; Synoeyenbos-West et al., 1999).

organism has only been recovered from miningimpacted lake sediments.

Geothrix Geothrix fermentans and closely related strains have been recovered from the Fe(III)-reducing zone of petroleum-contaminated aquifers (Anderson et al., 1998; Coates et al., 1999b). Like Geobacter species, G. fermentans can oxidize short-chain fatty acids to carbon dioxide with Fe(III) serving as the sole electron acceptor. It can also use long-chain fatty acids, as well hydrogen as an electron donor for Fe(III) reduction (Table 2) and can grow fermentatively on several organic acids. G. fermentans, along with Holophaga foetida, is part of a deeply branching group in the kingdom Acidobacterium. The 16S rDNA sequences from this kingdom are among the most common recovered from soil, but few organisms from this kingdom have been cultured (Barns et al., 1999). Studies in which Fe(III)-reducing microorganisms were recovered in culture media suggested that organisms closely related to G. fermentans might be as numerous as Geobacter species in the Fe(III) reduction zone of a petroleum-contaminated aquifer (Anderson et al., 1998). However, analyses of 16S rDNA sequences have indicated that Geothrix sp. are probably several orders of magnitude less numerous than Geobacter species in such environments (Rooney-Varga et al., 1999; Synoeyenbos-West et al., 1999).

Shewanella–Ferrimonas–Aeromonas In contrast to the organisms discussed above, which only grow anaerobically, several genera within the gProteobacteria, can grow aerobically, and under anaerobic conditions can use Fe(III), Mn(IV), or other electron acceptors (Table 2). These include species of Shewanella, Ferrimonas, and Aeromonas. Although many of these organisms can use a wide range of electron donors when oxygen is available as an electron acceptor, their range of electron donors with Fe(III) and Mn(IV) is generally restricted to hydrogen and small organic acids. An exception is Shewanella saccharophila, which also can use glucose as an electron donor for Fe(III) reduction. The Shewanella species, which have been studied in detail, incompletely oxidize multicarbon organic electron donors to acetate. Another Fe(III)-reducing microorganism that may be related to this group is an unidentified microorganism referred to as a “pseudomonad,” which was the first organism found to grow with hydrogen as the electron donor and Fe(III) as the electron acceptor (Balashova and Zavarzin, 1980). However, this organism does not appear to be available in culture collections for further study, and its true phylogenetic placement is unknown. The FMR in the g-Proteobacteria have been recovered from a variety of sedimentary environments including various aquatic sediments (Myers and Nealson, 1988; Caccavo et al., 1992; Coates et al., 1999a) and the subsurface (Pedersen et al., 1996; Fredrickson et al., 1998). However, in contrast to the organisms in the Geobacteraceae which are found to be numerous in both molecular and culturing analysis of widely diverse environments where Fe(III) reduction is important, the distribution of Shewanella is more variable. For example, Shewanella were found to account for ca. 2% of the microbial population in some surficial aquatic sediments, but could not be detected in other sediments (DiChristina and DeLong, 1993). Shewanella 16S rDNA sequences could not be recovered from aquifer sediments in which Fe(III) reduction was the predominant terminal electron-accepting process TEAP (Synoeyenbos-West et al., 1999). This was the case even when electron donors, such as lactate and formate, that are preferred by Shewanella species, were added to stimulate Fe(III) reduction.

Geovibrio ferrireducens and Deferribacter thermophilus Culturing from hydrocarbonimpacted soils and a petroleum reservoir have led to the recovery of the mesophile, Geovibrio ferrireducens (Caccavo et al., 1996) and the thermophile, Deferribacter thermophilus (Greene et al., 1997). These organisms are more closely related to each other than to any other known Fe(III)-reducing microorganisms and grow with similar electron donors for Fe(III)-reduction. G. ferrireducens has been shown to completely oxidize its carbon substrates to carbon dioxide and it is assumed that D. thermophilus can as well, but this has not been directly tested. An interesting feature of the metabolism of these organisms is the ability to use some amino acids as electron donors for Fe(III) reduction. The environmental distribution of these organisms has not been studied in detail. Ferribacter limneticum Ferribacter limneticum (Cummings et al., 1999) is the only organism in the b-subclass of the Proteobacteria that is known to conserve energy to support growth from Fe(III) reduction. Unlike many Fe(III)reducing microorganisms it does not utilize Mn(IV) as an electron acceptor. To date, this

Sulfurospirillum barnesii Sulfurospirillum barnesii which was initially isolated based on its ability to use selenate as an electron acceptor

646

D. Lovley

(Oremland et al., 1994b), also can grow using the reduction of Fe(III) and the metalloid As(V) (Laverman et al., 1995). Although it has commonly been found that if one organism in a close phylogenetic group has the ability to reduce Fe(III) then others in the group also will be Fe(III) reducers (Roden and Lovley, 1993a; Lovley et al., 1995c; Lonergan et al., 1996; Kashefi and Lovley, 1999), Sulfurospirillum arsenophilum does not reduce iron [Fe(III); Stolz et al., 1999)]. Wolinella succinogenes, which is also in the e-subclass of the Proteobacteria, also can reduce Fe(III) and metalloids (Lovley et al., 1997c; 1999b), but whether W. succinogenes conserves energy to support growth from metal reduction has not been determined. Acidophilic Fe(III)-Reducing Microorganisms Although Fe(III) is highly insoluble at the circumneutral pH at which most Fe(III)-reducing microorganisms have been studied, Fe(III) is soluble at low pH. The redox potential of the Fe+3/ Fe+2 redox couple is significantly more positive than the Fe(III) oxide/Fe+2 redox couple and the oxidation of electron donors (such as S∞) that might be unfavorable at circumneutral pH with Fe(III) oxides as the electron acceptors might be favorable in acidic pH where more Fe+3 is available. Thiobacillus ferroxidans can grow anaerobically with S∞ as the electron donor and Fe(III) as the electron acceptor (Das et al., 1992; Pronk et al., 1992). Thiobacillus thiooxidans also has been shown to reduce Fe(III) with S∞ as the electron donor (Brock and Gustafson, 1976), but the culture was grown aerobically and energy conservation from Fe(III) reduction was not demonstrated. This was also true of the thermophile, Sulfolobus acidocaldarius (Brock and Gustafson, 1976). Acidophilic thermophiles that can reduce Fe(III) with glycerol or thiosulfate as the electron donor have been described (Bridge and Johnson, 1998), but the ability of these organisms to conserve energy to support growth from Fe(III) reduction has not been examined in detail. An acidophilic mesophile, designated strain SJH, exhibited Fe(III)-dependent growth in a complex organic medium containing glucose and tryptone (Johnson and McGinness, 1991), but further characterization of the electron donors for Fe(III) reduction and a detailed description of the organism were not provided. Hyperthermophilic and Thermophilic Archaea and Bacteria In addition to D. thermophilus mentioned above, a number of other thermophiles and hyperthermophiles can conserve energy to support growth from Fe(III) reduction. The first thermophilic FMR reported was the deep subsurface isolate, Bacillus infernus, which

CHAPTER 1.21

has a temperature optimum of 60∞C (Boone et al., 1995). It was also the first Gram-positive FMR identified. In contrast to all other members of the Bacillus genus, B. infernus is a strict anaerobe and can grow by fermentation when Fe(III) or other electron acceptors are not available. Other thermophilic FMR recovered from subsurface environments include Thermoterrabacterium ferrireducens (Slobodkin et al., 1997) and a Thermus species (Kieft et al., 1999). As summarized in Tables 1 and 2, a wide phylogenetic diversity of hyperthermophilic microorganisms can transfer electrons to iron [Fe(III); Vargas et al., 1998)]. However, only three of these organisms, Pyrobaculum islandicum, P. aerophilum, and Thermotoga maritima, have been shown to conserve energy to support growth from Fe(III) reduction. P. islandicum and T. maritima grow with hydrogen as the electron donor and Fe(III) as the electron acceptor and P. islandicum and P. aerophilum also can grow with complex organic matter (peptone, yeast extract) as the electron donor and Fe(III) as the electron acceptor (Kashefi and Lovley, 1999). Forms of Fe(III) and Mn(IV) That Can Serve as Electron Acceptors Unlike other types of respiration that use soluble electron acceptors, Fe(III) and Mn(IV) reduction require the reduction of insoluble electron acceptors in most environments. The insoluble Fe(III) and Mn(IV) oxides that are the most environmentally relevant forms of Fe(III) and Mn(IV) at circumneutral pH can be found in a wide diversity of forms (Dixon and Skinner, 1992; Schwertmann and Fitzpatrick, 1992). The nature of the oxides have a major impact on the rate and extent of Fe(III) and Mn(IV) reduction (Lovley, 1991a; Lovley, 1995a). Pure cultures of Fe(III)-reducing microorganisms reduce a variety of insoluble Fe(III) and Mn(IV) forms (Lovley, 1991a), including the Fe(III) oxides naturally found in sedimentary environments (Lovley et al., 1990b; Coates et al., 1996). Early studies on Fe(III) reduction by fermentative microorganisms often employed highly crystalline Fe(III) oxides as the Fe(III) form (Table 1). However, studies on Fe(III) reduction in sediments suggested that the primary form of Fe(III) that FMR reduced in aquatic sediments was poorly crystalline Fe(III) oxides and that poorly crystalline Fe(III) oxides promoted the complete oxidation of organic compounds to carbon dioxide with Fe(III) serving as the electron acceptor (Lovley and Phillips, 1986a; Lovley and Phillips, 1986b; Phillips et al., 1993). The use of poorly crystalline Fe(III)-oxide as the Fe(III) form permitted the first recovery of

CHAPTER 1.21

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

a microorganism that could completely oxidize organic compounds to carbon dioxide with Fe(III) serving as the electron acceptor (Lovley et al., 1987c). Most subsequent studies that have enriched for Fe(III)-reducing microorganisms from the environment or that have evaluated mechanisms for Fe(III) oxide reduction by pure cultures of FMR have used poorly crystalline Fe(III) oxide as the electron acceptor. FMR have been shown to reduce some of the more crystalline Fe(III) oxides, including hematite, goethite, akaganeite, and magnetite, under some conditions (Table 2; Lovley, 1991a; Kostka and Nealson, 1995; Roden and Zachara, 1996). However, the rates of reduction of the crystalline Fe(III) oxides are generally much slower than the reduction of poorly crystalline Fe(III) oxide. In most instances, sustained growth is difficult to maintain in consecutive transfer of pure cultures with crystalline Fe(III) oxides as the electron acceptor. In evaluating the potential for reduction of crystalline Fe(III) oxides, it is important to omit complex organic matter or organic acids, which chelate and solubilize Fe(III) from the Fe(III) oxides. The FMR reduction of crystalline Fe(III) oxides in soils and sediments has not been demonstrated conclusively. An alternative, environmentally relevant, source of insoluble Fe(III) is structural Fe(III) in clays. Reduction of Fe(III) in clays is often observed in flooded soils and FMR have been shown to reduce this iron [Fe(III); Kostka et al., 1996; Lovley et al., 1998)]. Soluble Fe(III) forms are often used for culturing FMR. Although soluble Fe(III) may not represent an environmentally significant form of Fe(III), it provides an easy method for culturing FMR. Pure cultures generally reduce soluble Fe(III) forms faster than poorly crystalline Fe(III) oxide, and less insoluble precipitates are formed during reduction of soluble Fe(III). Furthermore, unlike poorly crystalline Fe(III) oxide, some soluble Fe(III) forms do not have to be synthesized because they are commercially available. Fe(III)-citrate is the most commonly used form of soluble Fe(III) for the culture of FMR. It is highly soluble and can readily be provided at concentrations as high as 50 mM, even in media with a high salt content. However, Fe(III)citrate may be toxic to some Fe(III)-reducing microorganisms (Lovley et al., 1990a; Lovley et al., 1993b; Roden and Lovley, 1993b). The Fe(III) chelated with nitrilotriacetic acid (Fe(III)-NTA) is a useful alternative. The limitations of Fe(III)-NTA are its frequent toxicity at concentrations above 10 mM and its tendency to precipitate as Fe(III) oxide when Fe(III)-NTA is added to media with high salt content or at temperatures of 60∞C or above. Unlike Fe(III)-

647

citrate, Fe(III)-NTA is not commercially available and must be synthesized, as described below. “Ferric pyrophosphate” has been successfully used for the culture of FMR (Caccavo et al., 1994; Caccavo et al., 1996). This is a somewhat undefined mixture that contains not only Fe(III) and phosphate, but also citrate and nitrilotriacetic acid which are likely to play an important role in maintaining the solubility of Fe(III) in this mixture. The most commonly used form of Mn(IV) oxide in studies of Mn(IV) reduction by FMR is birsnessite, a readily synthesized Mn(IV) oxide (see method for synthesis below). However, there is a wide diversity of Mn(IV) oxides is found in the environment and rates of Mn(IV) reduction can be dependent upon the form of Mn(IV) oxide available (Burdige et al., 1992). Products of Fe(III) and Mn(IV) Reduction Products Fe(II) and Mn(II) are more soluble than Fe(III) and Mn(IV) and thus microbial Fe(III) and Mn(IV) reduction results in a marked increase in dissolved iron and manganese in anaerobic environments and in cultures of FMR. However, in both cultures and sediments, most of the Fe(II) and Mn(II) produced during microbial reduction of insoluble Fe(III) and Mn(IV) oxides often remains in solid forms (Lovley, 1991a; Lovley, 1995a; Schnell et al., 1998). In culture, microbial Fe(III) and Mn(IV) reduction has been shown to form such minerals as magnetite (Fe3O4) siderite (FeCO3), vivianite (Fe3PO4 · 8H 2O) and rhodochrosite (MnCO3; Lovley, 1991a; Lovley, 1995b). The formation of such minerals in culture provides a model for the geologically significant deposition of iron and manganese minerals described above. The fact that most of the Fe(II) and Mn(II) produced from microbial Fe(III) and Mn(IV) reduction is insoluble means that quantitative analysis of Fe(III) or Mn(IV) reduction either in cultures or environmental samples requires quantifying the amount of insoluble Fe(II) or Mn(II) produced. The Fe(II) may be solubilized in HCl (Lovley and Phillips, 1986a) or oxalate (Phillips and Lovley, 1987; Lovley and Phillips, 1988c) before measurement with Fe(II)-specific reagents such as ferrozine (Stookey, 1970) or ion chromatography (Schnell et al., 1998). Loss of Fe(III) in acid-solubilized samples also can be monitored (Lovley and Phillips, 1988b; Schnell et al., 1998). Methods for quantitatively measuring Mn(IV) reduction are not as well established. Much of the Mn(II) produced during Mn(IV) reduction adsorbs onto the Mn(IV) oxide or forms insoluble Mn(II) minerals. Mn(II) can be solubilized in acid and soluble manganese measured with atomic absorption spectroscopy (Lovley and

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D. Lovley

Phillips, 1988c), but this is technically difficult because acid will also eventually dissolve the Mn(IV) oxide. A better strategy might be to solubilize all the manganese and specifically measure the Mn(II) produced with ion chromatography (Schnell et al., 1998). Mechanisms for Electron Transfer to Fe(III) and Mn(IV) The mechanisms by which Fe(III)and Mn(IV)-reducing microorganisms transfer electrons to insoluble Fe(III) and Mn(IV) are poorly understood. It is generally stated that Fe(III) and Mn(IV) reducers must directly reduce Fe(III) and Mn(IV) oxides by establishing contact with the oxides (Lovley, 1991a). Until recently, the primary evidence of the need for contact was the finding that Fe(III) and Mn(IV) were not reduced when Fe(III) or Mn(IV) oxides and Fe(III)- and Mn(IV)-reducing microorganisms were separated by semipermeable membranes, which should permit the passage of soluble substances. This result as well was considered evidence that Fe(III)- and Mn(IV)reducing microorganisms do not produce chelators to solubilize Fe(III) or Mn(IV) and do not produce compounds that could serve as soluble electron-shuttles between Fe(III)- and Mn(IV)reducing microorganisms and the insoluble oxides. However, recent studies have demonstrated that this approach is flawed because even when chelators or electron shuttles were added to cultures, Fe(III)-reducing microorganisms still did not significantly reduce Fe(III) oxide held within dialysis tubing (Nevin and Lovley, 1999a). Studies with strains of Shewanella alga, which were deficient in the ability to attach to Fe(III) oxides, continued to reduce Fe(III), suggesting that attachment to Fe(III) oxide was not necessary for Fe(III) oxide reduction (Caccavo et al., 1997). Thus, although studies have documented the association of Fe(III)-reducing microorganisms with Fe(III)-oxide particles, the current evidence is not definitive to clearly state that Fe(III)- and Mn(IV)-reducing microorganisms must attach to Fe(III) and Mn(IV) oxides in order to reduce them. It was suggested that Geobacter sulfurreducens might reduce Fe(III) oxide in culture by releasing a low molecular weight (9.6 kDa) c-type cytochrome into the medium which could serve as a soluble electron shuttle between G. sulfurreducens and the Fe(III) oxide (Seeliger et al., 1998). However, further investigation has demonstrated that this c-type cytochrome is not an effective electron shuttle and that in healthy, actively growing cultures of G. sulfurreducens, little, if any, of the 9.6 kDa cytochrome is released into the growth medium (Lloyd et al., 1999). Therefore, the proposed shuttling mechanism is unlikely.

CHAPTER 1.21

Iron [Fe(III)]-reducing microorganisms can use humics and other extracellular quinones as electron shuttles to promote Fe(III) oxide reduction (Lovley et al., 1996; Lovley et al., 1998; Lovley et al. 2000). As discussed below, humics and other extracellular quinones can serve as electron acceptors for Fe(III)-reducing microorganisms. The hydroquinone moieties that are generated as the result of the reduction of extracellular quinones can transfer electrons to Fe(III) oxides through a strictly abiotic reaction. This reduction of Fe(III) regenerates quinone moieties that can then again serve as electron acceptors for Fe(III)-reducing microorganisms. In this manner a small amount of extracellular quinone can promote a significant increase in the rate of reduction of poorly crystalline Fe(III) oxide. For example, studies with cultures and aquifer sediments have demonstrated that there is a significant potential for electron shuttling with as little as 100 nM AQDS (Lloyd et al., 1999; Nevin and Lovley, 1999b). Although electron shuttling to Mn(IV) oxides have not been studied in detail, a similar phenomenon is expected. However, both the evidence that Fe(III)- and Mn(IV)-reducing microorganisms can reduce Fe(III) and Mn(IV) oxides in cultures without added electron shuttling compounds and chelators and the lack of evidence for release of electron shuttling or chelating compounds by the microorganisms (Nevin and Lovley, 1999a) suggests that FMR can directly transfer electrons to Fe(III) and Mn(IV) oxides. The Fe(III)reductase activity is primarily localized in the membranes of Fe(III)- and Mn(IV)-reducing microorganisms such as G. metallireducens (Gorby and Lovley, 1991), S. putrefaciens (Myers and Myers, 1993), and G. sulfurreducens (Gaspard et al., 1998; Magnuson et al., 1999). The involvement of cytochromes of the c-type has been suggested to be involved in electron transport to Fe(III) in G. metallireducens (Lovley et al., 1993c) and S. putrefaciens (Myers and Myers, 1992; Myers and Myers, 1997; Beliaev and Saffarini, 1998). A NADH-dependent Fe(III) reductase complex was purified from G. sulfurreducens and a 90-kDa c-type cytochrome in the complex served as the Fe(III) reductase (Magnuson et al., 1999). However, no study has as yet definitively identified as yet the physiologically relevant Fe(III) or Mn(IV) reductase in any organism capable of conserving energy to support growth via Fe(III) or Mn(IV) reduction.

Other Respiratory Capabilities of FMR Many FMR can reduce other electron acceptors well-known to support anaerobic respiration such as fumarate, nitrate, and S∞ (Table 2). Fumarate is reduced to succinate, and S∞ is

CHAPTER 1.21

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

reduced to sulfide. In those documented instances of nitrate reduction, nitrite or ammonia has been found to be the product. It is interesting that nearly all microorganisms with the ability to reduce Fe(III) also can reduce S∞ to sulfide. In fact, screening of known S∞-reducing microorganisms already available in culture has been a fruitful approach for discovering new FMR (Roden and Lovley, 1993a; Lonergan et al., 1996; Vargas et al., 1998). Electron Transfer to Other Metals and Metalloids Many Fe(III)-reducing microorganisms can transfer electrons to metals other than iron or manganese [Fe(III) or Mn(IV); Table 2]. For example, G. metallireducens and S. putrefaciens can grow with U(VI) as the sole electron acceptor (Lovley et al., 1991b). Cell suspensions of other FMR have been found to transfer electrons to U(VI), but their ability to obtain energy to support growth from U(VI) reduction has not been evaluated. Many sulfatereducing microorganisms, can effectively reduce U(VI), but attempts to grow these organisms with U(VI) as the sole electron acceptor have been unsuccessful (Lovley et al., 1993b). U(VI), which is soluble in bicarbonate-based media is reduced to U(IV) that precipitates as the mineral uraninite (Gorby and Lovley, 1992; Lovley and Phillips, 1992). Visualization of microbial U(VI) reduction can be enhanced with the use a fluorescent light. The U(VI)-containing liquid cultures or agar plates fluoresce green, whereas the uraninite does not significantly fluoresce. Loss of U(VI) during U(VI) reduction can be monitored as loss of soluble uranium by monitoring total uranium concentrations in culture filtrates, but since U(IV) precipitation is not instantaneous (Gorby and Lovley, 1992), more quantitative estimates of U(VI) reduction can be more quantitatively estimated by monitoring loss of U(VI) with a kinetic phosphorescence analyzer (Lovley et al., 1991b) or by using ion chromatography. Several Fe(III)-reducing microorganisms can reduce the oxidized form of the radioactive metal technetium, Tc(VII) to reduced forms (Table 2). Growth with Tc(VII) as the sole electron acceptor has not yet been documented as yet in any organism. Tc(VII) reduction can be monitored by following the formation of reduced technetium forms with paper chromatography and a phosphorimager (Lloyd and Macaskie, 1996). FMR can reduce a variety of other metals and metalloids (Table 2). Several can reduce Cr(VI) to Cr(III), but growth with Cr(VI) as the sole electron acceptor has not been demonstrated (Lovley, 1995c). The FMR, S. barnesii can conserve energy from the reduction of Se(VI) to

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Se∞ and As(V) to As(III) (Laverman et al., 1995). Electron Transfer to and from Humic Substances and Other Extracellular Quinones All FMR that have been evaluated to date, including the hyperthermophiles, have the ability to transfer electrons to humic substances (humics) or other extracellular quinones such as the humics analog, anthraquinone-2,6-disulfonate (AQDS) Lovley et al., 1996; Lovley et al., 1998; Lovley et al., 2000). In those organisms in which the potential for growth has been evaluated, energy to support growth is from electron transport to humics and this capability is conserved. Electron-spin resonance (ESR) studies have suggested that quinones are important electron-accepting groups in the humics (Scott et al., 1998). The ESR studies with AQDS as the sole electron acceptor have directly demonstrated that energy can be conserved from electron transfer to extracellular quinones has been directly demonstrated in studies with AQDS as the sole electron acceptor (Lovley et al., 1996; Coates et al., 1998; Lovley et al., 1998). Humics can chelate Fe(III) that is also available for microbial reduction (Benz et al., 1998; Lovley and Blunt-Harris, 1999a), but the concentration of microbially reducible Fe(III) in humics is a minor fraction of the total electron-accepting capacity (Lovley and Blunt-Harris, 1999a). A wide diversity of humics can serve as electron acceptors for Fe(III)-reducing microorganisms (Lovley et al., 1996; Scott et al., 1998). Highly purified reference humics that have been extracted from diverse environments can be obtained from the International Humic Substances Society. Other commercially available humics are highly impure, differ from humics found in soils and sediments, and therefore should be avoided for definitive studies because commercially available humics are highly impure and their characteristics are unlike the humics found in soils and sediments (Malcolm and MacCarthy, 1986). The expense and technical difficulty of conducting studies with humics makes it preferable to carry out some studies on microbial reduction of extracellular quinones with humics analogs, such as AQDS (Lovley et al., 1996; Lovley et al., 1998). The advantages of AQDS are its low cost, high solubility, and its easy detection [an orange color develops when AQDS is reduced to anthrahydroquinone-2,6-disulfonate (AHQDS)]. Several FMR have the ability to use reduced extracellular quinones as an electron donor for reduction of electron acceptors such as nitrate and fumarate (Lovley et al., 1999b). Shewanella alga and Geobacter sulfurreducens grew with

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D. Lovley

AHQDS as the electron donor. However, other FMR that could oxidize AHQDS in cell suspensions could not be grown with AHQDS as the sole electron acceptor. The ability of FMR to both reduce and oxidize extracellular quinones permits their use with other quinone-oxidizing and quinone-reducing microorganisms as an interspecies electron transfer system in which quinones serve as the electron shuttle between the microorganisms (Lovley et al., 1999b). Proton Reduction in Syntrophic Association with Hydrogen-Consuming Microorganisms In the absence of Fe(III) or other suitable electron acceptors, some organisms in the Geobacteraceae can transfer electrons to protons to produce hydrogen gas. For hydrogen production to be thermodynamically favorable, a sink for hydrogen, such as a hydrogen-consuming microorganism, must keep hydrogen concentrations low enough. For example, several Pelobacter species can oxidize ethanol to acetate and carbon dioxide when grown in association with hydrogen-consuming microorganisms (Schink, 1992). G. sulfurreducens can oxidize acetate to carbon dioxide when cultured with Wolinella succinogenes, which oxidizes hydrogen with concomitant reduction of nitrate (Cord-Ruwisch et al., 1998). Reductive Dechlorination Several Fe(III)reducing microorganisms are capable of using chlorinated compounds as electron acceptors. Desulfuromonas chlorethenica, which was isolated as a tetrachloroethylene-respiring microorganism (Krumholz et al., 1996; Krumholz, 1997), was found to grow also with Fe(III) as the electron acceptor, as expected for microorganisms within the family Geobacteraceae (Lonergan et al., 1996). Other Geobacteraceae that were evaluated did not reduce tetrachloroethylene. Desulfitobacterium dehalogenans which can use chlorophenolic compounds as an electron acceptor (Utkin et al., 1994), also can grow with Fe(III) as the electron acceptor (Lovley et al., 1998). Another chlorophenol-respiring species in the same genus, Desulfitobacterium hafniense, was reported to reduce Fe(III), but it was not reported whether growth was conserved from Fe(III) reduction (Christiansen and Ahring, 1996).

Recovery of Fe(III)- and Mn(IV)-Reducing Microorganisms in Culture Localizing Zones of Fe(III) and Mn(IV) Reduction Although FMR can be recovered from nearly any soil or sediment sample, it is generally of interest to study organisms from

CHAPTER 1.21

habitats in which Fe(III) and Mn(IV) are ongoing processes. Dissimilatory Fe(III) and Mn(IV) reduction are geochemically most significant in anaerobic environments such as freshwater and marine sediments; flooded soils or the anaerobic interior of soil aggregates; the deep terrestrial subsurface; and shallow aquifers contaminated with organic compounds. In aquatic sediments and the terrestrial subsurface Fe(III) and Mn(IV) reduction are most apparent in discrete anoxic sediment layers in which the endproducts of Fe(III) and Mn(IV) reduction, Fe(II) or Mn(II), are accumulating. In the typical zonation of respiratory processes found with depth in aquatic sediments or along the groundwater flow path in the subsurface, the zones of Fe(III) and Mn(IV) reduction are typically bounded on one side by the zone of nitrate reduction and on the other side by the zone of sulfate reduction (Lovley and Chapelle, 1995c). In addition to these larger discrete zones of Fe(III) reduction and Mn(IV) reduction in sedimentary environments, it is important to recognize that many soils and sediments that are predominately aerobic alsomay contain abundant anaerobic microzones in which Fe(III) and Mn(IV) reduction may be taking place. Although accumulation of dissolved Fe(II) and Mn(II) in groundwater or porewater can be used to help identify the zones of Fe(III) and Mn(IV) reduction in subsurface or aquatic sediments, such standard geochemical measurements can often fail to accurately locate the metal reduction zones (Lovley et al., 1994b). A primary reason for this failure is that high concentrations of Fe(II) and Mn(II) may be found in sediments in which other TEAPs, such as methanogenesis, predominate. In environments where conditions approach steady-state, such as aquatic sediments and aquifers, measurements of dissolved hydrogen can be used to identify zones in which Fe(III) reduction is the TEAP (Lovley and Goodwin, 1988a; Lovley et al., 1994c). This is because there is a unique range of dissolved hydrogen that is associated with Fe(III) reduction that is the predominant TEAP in steady-state environments. Hydrogen measurements have not been used to localize Mn(IV)-reducing zones because: 1) hydrogen concentrations under Mn(IV)-reducing conditions are very low and difficult to accurately measure accurately; 2) hydrogen concentrations for Mn(IV) and nitrate reduction are similar; and 3) the low concentrations of Mn(IV) in many soils means that the Mn(IV) reduction zone is not extensive. An alternative method for determining the zone of Fe(III) reduction in soils and sediments is to use [2-14C]-acetate (Lovley, 1997a). The reduction of Fe(III) can be considered to be the

CHAPTER 1.21

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

TEAP if: 1) a tracer quantity of [2-14C]-acetate added to the sediments is converted to 14CO2 with no production of 14CH4; 2) the production of 14CO2 is not inhibited with the addition of molybdate; 3) the sediments are depleted of nitrate; and 4) the sediments contain some Fe(II). The reasoning for this is that: 1) lack of 14 CH4 production rules out methanogenesis as a TEAP; 2) molybdate inhibits acetate oxidation by sulfate reducers so the lack of inhibition with molybdate rules out sulfate reduction as the TEAP; 3) nitrate reduction can not be an important TEAP in the absence of nitrate; and 4) Mn(IV) reduction can not be the TEAP in the presence of Fe(II) because Fe(II) rapidly reacts with Mn(IV) (Lovley and Phillips, 1988b) and thus Fe(II) will only be found if reactive Mn(IV) has been depleted. The rates of other TEAPs can often be quantified in sediments with the use of radiotracers. Unfortunately, attempts to measure rates of Fe(III) reduction in sediments with radioactively labeled Fe(III) were unsuccessful (Roden and Lovley, 1993b). This was because there was rapid isotope exchange between the radiolabelled Fe(III) and other iron pools, including Fe(II), was rapid. Thus, it was not possible to monitor rates of microbial Fe(III) reduction by measuring the production of radiolabeled Fe(II) from labeled Fe(III). Rates of Fe(III) and Mn(IV) reduction in sediments can be estimated from anaerobic incubations of sediments by monitoring the accumulation of Fe(II) and Mn(II) are monitored over time. It is important that the solid phase Fe(II) and Mn(II) pools be measured after acidic extractions or some other technique because most of the Fe(II) and Mn(II) are not recovered in the dissolved phase (Lovley and Phillips, 1988c; Lovley, 1991a). Geochemical modeling has been used to estimate rates of Fe(III) and Mn(IV) reduction in some aquatic sediments and subsurface environments and potentially could be used to identify zones of Fe(III) and Mn(IV) reduction (Lovley, 1995a). Isolation Procedures Although some FMR also can use oxygen as an electron acceptor or are tolerant of exposure to air, many are strict anaerobes. Therefore, unless the goal is to specifically select for facultative FMR, the use of strict anaerobic technique is preferable in initial enrichment and/or isolation procedures. To date, most FMR have been recovered using slight modifications of standard (Miller and Wolin, 1974; Balch et al., 1979) anaerobic techniques. This involves the use of culture tubes or bottles fitted with thick butyl rubber stoppers; removing traces of oxygen from gases by passing the gases through a column of heated copper filings; and

651

carrying out transfers with syringes and needles or under a stream of anoxic gas. Culture media can be prepared with the classical approach (Hungate, 1969) of boiling the media under a stream of anoxic gas to remove dissolved oxygen and then dispensing into tubes or bottles under anaerobic conditions. Alternatively, aerobic media may be dispensed into individual tubes or bottles and then the media can be vigorously bubbled with anoxic gas to strip dissolved oxygen from the media (Lovley and Phillips, 1988c). Both media preparation approaches appear to yield similar organisms. Reducing agents such as Fe(II)—typically supplied at 1–3 mM as ferrous chloride—cysteine (0.25–1 mM), or sulfide (0.25–1 mM) can be added to dispensed media from anoxic stocks just prior to inoculation. In addition to reacting with any trace oxygen in the media, cysteine and sulfide will reduce Fe(III) and Mn(IV) in the media, producing Fe(II) and Mn(II). Fe(II) rapidly reacts with traces of oxygen, forming Fe(III). Manganese [Mn(II)] will only slowly react abiotically with oxygen. Many FMR have been recovered without the addition of reducing agents to the media. Once Fe(III) reduction begins, the Fe(II) formed serves as protection against oxygen contamination. Reducing agents are rarely used in media designed for liquid-to-liquid transfer of Fe(III)-reducing cultures because the inoculum of the Fe(III)-reducing cultures typically contain millimolar quantities of dissolved Fe(II), which will scavenge traces of oxygen from the media to which the inoculum has been added. A variety of media has been successfully employed for the enrichment and isolation of FMR, many of which are given in the references provided with each of the organisms in Table 2. An example of a freshwater and a marine medium are provided below. No definitive comparative studies of the efficacy of various media in recovering FMR have been carried out. However, it has been found that the freshwater medium described here can be used to recover Geobacter species with 16S rDNA sequences that are closely related to the 16S rDNA sequences that predominate in the Fe(III) reduction zone of sandy aquifers (Rooney-Varga et al., 1999; Synoeyenbos-West et al., 1999). Most successful isolations of pure cultures of Fe(III)- and Mn(IV)-reducing microorganisms have used either organic acids, primarily acetate or lactate, or hydrogen as the electron donor. If an enrichment step is used in the initial stages of recovery of the organisms, then fermentable compounds such as glucose generally result in the enrichment of fermentative microorganisms. However, as summarized above, some Fe(III)and Mn(IV)-reducing microorganisms can use sugars and amino acids as electron donors and

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these electron donors potentially could be be used for direct isolation of FMR. A variety of Fe(III) and Mn(IV) forms that were discussed above can be used as electron acceptors for enrichment or isolation. Iron added as Fe(III)-citrate and Fe(III) pyrophosphate is not ideal for enrichment cultures as the citrate is rapidly degraded by microorganisms other than Fe(III) reducers. Once the citrate is degraded, the Fe(III) from the Fe(III)-citrate precipitates as an insoluble Fe(III) oxide and thus defeats the purpose of adding the chelator. The compound Fe(III)NTA is relatively resistant to anaerobic degradation and can be used as a soluble source of Fe(III) for enrichment of Fe(III) reducers. However, as noted above, it is not suitable for use in media with marine salinities or at high temperature. Both Fe(III)-citrate and Fe(III)NTA are toxic to some Fe(III) reducers. Although solubilization of Mn(IV) with various chelators for use in recovery of Mn(IV)reducing microorganisms may be possible, this approach has not been widely used. As noted above, poorly crystalline Fe(III) oxide is typically the insoluble Fe(III) oxide of choice for culturing. A wide diversity of other Fe(III) oxides can be synthesized (Schwertmann and Cornell, 1991), if desired. If the media is dispensed aerobically into culture vessels, then a slurry of the Fe(III) or Mn(IV) oxide can be added to the vessels prior to addition of the media. An advantage of using poorly crystalline Fe(III) oxide as the electron acceptor is that most Fe(III)-reducing microorganisms convert the poorly crystalline Fe(III) oxide to the magnetic mineral magnetite during reduction. This is visually apparent as the reddish, non-magnetic Fe(III) oxide is transformed into a black, highly magnetic precipitate (Lovley et al., 1987c). Reduction of the Mn(IV) oxide is also visually apparent in bicarbonate-buffered media because reduction of the dark Mn(IV) oxide results in its dissolution and concomitant accumulation of rhodochrosite, a white Mn(II) carbonate mineral. An alternative electron acceptor that can be used for the recovery of Fe(III)- and Mn(IV)reducing microorganisms is the humics analog, AQDS, which is typically provided at 5 mM. All of the Fe(III)-reducing microorganisms that have been evaluated can reduce AQDS, whereas microorganisms that do not reduce Fe(III) can not reduce AQDS (Lovley et al., 1996; Lovley et al., 1998; Lovley et al., 2000). Recovery of AQDS-reducing microorganisms either through enrichment and isolation procedures or dilutionto-extinction approaches yield organisms that also can reduce iron [Fe(III); Coates et al., 1998)]. The reduction of AQDS to AHQDS is

CHAPTER 1.21

visually apparent as the conversion of the relatively colorless AQDS to the orange, AHQDS. Fe(III)- and Mn(IV)-reducing microorganisms can be obtained in pure culture through standard anaerobic approaches of isolating colonies in tubes or on plates or through dilution-toextinction in liquid media. Soluble Fe(III) forms or AQDS are often used for isolating colonies on agar-solidified media, but colonies also can be obtained by incorporating Fe(III) and Mn(IV) oxides into solidified media. The Fe(III)- and Mn(IV)-reducing microorganisms that have the ability to use other electron acceptors often can be successfully purified from Fe(III)- or Mn(IV)reducing enrichment cultures with these alternative electron acceptors. Common alternative electron acceptors include nitrate, fumarate, sulfur, and oxygen. Suggested Media for Enrichment and Culturing of FMR Freshwater and marine media suitable for culturing a diversity of mesophilic FMR are described below. A variety of other media have also been used which can be found in the references for the individual organisms. The media described here have a bicarbonate-carbon dioxide buffer system and the headspace gas typically contains 20% carbon dioxide to establish an initial pH of ca. 6.8. Freshwater Medium To 900 ml water add: NaHCO3 NH4Cl NaH2PO4 · H 2O KCl Vitamin Solution Mineral Solution

2.50 g 0.25 g 0.60 g 0.10 g 10.0 ml 10.0 ml

Bring solution to a final volume of 1 liter. Media is dispensed, sparged with an 80:20 mixture of N2:CO2 gas and then autoclaved.

Marine Medium Medium contains, per liter: NaCl KCl NaHCO3 Vitamin solution Mineral solution RST minerals stock Salt stock*

20.0 g 0.67 g 2.5.0 g 10.0 ml 10.0 ml 20.0 ml 50.0 ml

*Add salt solution aseptically and anaerobically after autoclaving.

RST Minerals Stock 50X Stock contains, per 100 ml: NH4Cl KCl KH2PO4 MgSO4 · 7H 2O CaCl2 · 2H 2O

5.0 g 0.5 g 0.5 g 1.0 g 0.1 g

CHAPTER 1.21

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

citrate [typically 13.7 g to provide a final concentration of ca. 50 mM Fe(III)]. Once the ferric citrate is dissolved quickly cool the medium to room temperature in an ice bath. Adjust pH to 6.0 using 10N NaOH. When the pH approaches 5.0, add the NaOH dropwise. Add medium constituents as outlined above. Bring to a final volume of 1 liter. Do not expose this media to direct sunlight to prevent photoreduction of the Fe(III).

Salt Stock Stock contains, per 100 ml: MgCl2 · 6H 2O CaCl2 · 2H 2O

21.2 g 3.04 g

Vitamin Solution Solution contains, per liter: Biotin Folic acid Pyridoxine HCl Riboflavin Thiamine Nicotinic acid Pantothenic acid B-12 p-Aminobenzoic acid Thioctic acid

2.0 mg 2.0 mg 10.0 mg 5.0 mg 5.0 mg 5.0 mg 5.0 mg 0.1 mg 5.0 mg 5.0 mg

Mineral Solution grams per liter Trisodium nitrilotriacetic acid MgSO4 MnSO4 · H 2O NaCl FeSO4 · 7H 2O CaCl2 · 2H 2O CoCl2 · 6H 2O ZnCl2 CuSO4 · 5H 2O AlK(SO4)2 · 12H 2O H3BO3 Na2MoO4 NiCl2 · 6H 2O Na2WO4 · 2H 2O

653

1.5 g 3.0 g 0.5 g 1.0 g 0.1 g 0.1 g 0.1 g 0.13 g 0.01 g 0.01 g 0.01 g 0.025 g 0.024 g 0.025 g

Preparation of Fe(III) and Mn(IV) Forms Poorly Crystalline Fe(III) Oxide Dissolve FeCl3·6H 2O in water to provide final concentration of 0.4M. Stir continually while SLOWLY adjusting the pH to 7.0 dropwise with 10 M NaOH solution. It is extremely important not to let the pH rise above pH 7 even momentarily during the neutralization step because this will result in an Fe(III) oxide that is much less available for microbial reduction. Continue to stir for 30 minutes once pH 7 is reached and recheck pH to be sure it has stabilized at pH 7. To remove dissolved chloride, centrifuge the suspension at 5,000 rpm for 15 minutes. Discard the supernatant, resuspend the Fe(III) oxide in water, and centrifuge. Repeat six times. On the last wash, resuspend the Fe(III) oxide to a final volume of approximately 400 ml, and after determining iron content, adjust Fe(III) concentration to approximately 1 mole per liter. Typically, Fe(III) oxide is added to individual tubes of media to provide 100 mmol per liter. Fe(III)-Citrate Prior to the addition of any of the media constituents, heat 800 ml of water on a stirring hot-plate to near boiling. Add Fe(III)-

Fe(III) Nitrilotriacetic Acid To make a stock of 100 mM Fe(III)-NTA, dissolve 1.64 g of NaHCO3 in 80 ml water. Add 2.56 g C6H6NO6Na3 (sodium nitrilotriacetic acid) and then 2.7 g FeCl3·6H 2O. Bring the solution up to 100 ml. Sparge the solution with N2 gas and filter sterilize into a sterile, anaerobic serum bottle. Do not autoclave. Typically, 100 mM Fe(III)-NTA stock is added to individual tubes of media to provide a final concentration of 5 or 10 mmol of Fe(III). Goethite Prepare a 0.4M FeCl3·6H 2O solution. With continual stirring, adjust the pH to between 11 and 12 with 10 M NaOH solution. The suspension will become very thick. Ensure continual stirring and rinse the pH electrode frequently. The color of this suspension will turn to an ochre color as goethite is formed. One week at room temperature followed by 16 hours at 90∞C is sufficient to convert the Fe(III) to goethite. The suspension should be washed to remove chloride, as described above for poorly crystalline Fe(III) oxide. The formation of goethite should be confirmed by X-ray diffraction analysis. The Fe(III) oxide also should be tested with extractants (Lovley and Phillips, 1987b; Phillips and Lovley, 1987) to ensure that it does not contain poorly crystalline Fe(III) oxide. Hematite Hematite is readily available from chemical supply companies as “Ferric Oxide.” Manganese Oxide To 1 liter of a solution containing 80 mM NaOH and 20 mM KMnO4 slowly add 1 liter of 30 mM MnCl2 with mixing. Wash the manganese oxide precipitate, as described above for poorly crystalline Fe(III) oxide, to lower the dissolved chloride concentration. Enumeration of Fe(III)- and Mn(IV)-Reducing Microorganisms The FMR in environments can be enumerated with standard most-probablenumber (MPN) culturing techniques using variations of media described above. Enumerations typically use Fe(III) or AQDS as the electron acceptor with the understanding that the Fe(III)reducing microorganisms recovered are likely to

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have the ability to reduce Mn(IV) as well. Poorly crystalline Fe(III) oxide or Fe(III)-NTA is preferred over Fe(III)-citrate and Fe(III)pyrophosphate, which promote the growth of fermentative microorganisms. One successful approach has been to add a combination of poorly crystalline Fe(III) oxide (100 mmol/liter) and 4 mM NTA to provide a supply of chelated Fe(III). FMR also can be counted in plate counts in which Fe(III)-NTA or AQDS has been added as the electron acceptor. Clearing zones develop around FMR reducing Fe(III)-NTA, and growth with AQDS as the electron acceptor results in the formation or orange colonies or zones. When possible, molecular enumeration rather than viable culturing enumeration techniques are the preferred methods because of the potential biases associated with the latter. The wide phylogenetic diversity of dissimilatory Fe(III) reducing microorganisms and the lack of an identified conserved gene associated with Fe(III) reduction make it impossible to enumerate Fe(III)-reducing microorganisms with one specific gene sequence (Lonergan et al., 1996). However, target 16S rRNA sequences that are selective for known groups of Fe(III)-reducing microorganisms have been identified and have been used to study the distribution of Fe(III)reducing microorganisms in sedimentary environments (DiChristina and DeLong, 1993; Anderson et al., 1998; Rooney-Varga et al., 1999; Synoeyenbos-West et al., 1999).

Summary Microbial reduction of Fe(III) and Mn(IV) is of environmental significance in a variety of aquatic sediments and the subsurface, influencing both the carbon cycle and the fate of many metals and metalloids, in both pristine and contaminated environments. Geological and microbiological evidence suggests that Fe(III) reduction was one of the earliest forms of respiration. A wide phylogenetic diversity of Fe(III)- and Mn(IV)reducing microorganisms have been recovered in pure culture, but with the exception of the recently recognized importance of Geobacter in subsurface environments, little is known about the distribution or relative contributions of the various Fe(III)-reducing microorganisms. The study of the mechanisms of Fe(III) and Mn(IV) reduction are also in their infancy. However, now that methods for culturing these organisms are well developed, it seems likely that increased insight into the ecophysiology of Fe(III)- and Mn(IV)-reducing microorganisms is forthcoming.

CHAPTER 1.21

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Prokaryotes (2006) 2:659–768 DOI: 10.1007/0-387-30742-7_22

CHAPTER 1.22 yrot a l imi s s iD

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Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes RALF RABUS, THEO A. HANSEN AND FRIEDRICH WIDDEL

Introduction A unique characteristic in the prokaryotic world is the multiplicity of life strategies without any involvement of oxygen. Actually, life in anoxic habitats is prokaryotic to a large extent. Prokaryotes have evolved not only various fermentation pathways, but also the capacity to couple the oxidation of organic substrates to the reduction of inorganic compounds (other than O2) to conserve energy for anaerobic growth. Electron acceptors reduced by prokaryotes under anoxic conditions are nitrate, manganese(IV), ferric iron, sulfate, elemental sulfur, other sulfur species (e.g., thiosulfate), carbon dioxide, protons and even oxidized forms of naturally less abundant elements such as arsenate(V), chromate(VI), selenate and uranium(VI). In several prokaryotes, even the electron donor may be inorganic, which results in purely inorganic (lithotrophic) redox reactions for energy conservation under anoxic conditions; notable examples are the oxidation of sulfur species with nitrate, or of molecular hydrogen with nitrate, iron(III), sulfate, sulfur or CO2. Two organic compounds with some relationship to inorganic electron acceptors are dimethylsulfoxide (DMSO) and trimethylamine-N-oxide (TMAO). In these compounds, anaerobic microorganisms reduce the oxygenated sulfur or nitrogen moiety, respectively. In most cases, the electron transport to the inorganic electron acceptors is associated with a mode of energy conservation that may be regarded as an anaerobic analogue to respiration with O2. This is particularly evident if the only electron donor is H2. In such a process, ATP synthesis can be only explained by a chemiosmotic transmembrane process rather than by fermentative substrate-level phosphorylation. Because of this analogy to the known respiratory chain, growth by utilization of inorganic electron acceptors other than O2 is usually termed “anaerobic respiration”. In some microorganisms, inorganic compounds (as for instance ferric iron or sulfur) may be reduced in by-reactions without obvious connection to respiration-like chemiosmotic energy conservation. Such by-

reactions may facilitate fermentation (disposal of reducing equivalents) but they should not be termed “anaerobic respirations”. Interestingly, most types of anaerobic respirations have not been encountered so far in the eukaryotic domain. The only (thus far reported) case of anaerobic respiration in a eukaryote is nitratereduction by a flagellate (Finlay et al., 1983). The microbial reduction of inorganic compounds contributes significantly to the global cycling of elements and represents the counterpart to oxidative microbial processes, e.g., nitrification, iron oxidation and sulfur oxidation. Among the anaerobic respirations, the reduction of sulfur species is most striking because it gives rise to a conspicuous end product, hydrogen sulfide (H2S), which is commonly known as a toxic chemical with a characteristic smell. By its chemical reactivity (e.g., toward iron minerals and oxygen), H2S has a pronounced impact on the chemistry of the environment. Despite of its toxicity, sulfide serves as electron donor for a great diversity of aerobic chemotrophic and anoxygenic phototrophic microorganisms that may form visible blooms in sulfidic habitats. The natural reduction and oxidation of sulfur species is known as the sulfur cycle. Because sulfate is the thermodynamically stable and most abundant form of sulfur in our oxic biosphere, sulfate reduction forms the basis of the biological sulfur cycle (Henrichs and Reeburgh, 1987; Jørgensen, 1987; Skyring, 1987; Widdel, 1988). A great diversity of sulfate-reducing microorganisms has been isolated from aquatic habitats. The chemical and biological oxidation processes of sulfide do not always lead directly to sulfate, but often yield intermediate oxidation states such as elemental sulfur or thiosulfate. These may serve as electron acceptors for anaerobic microorganisms that cannot reduce sulfate. Among these, sulfur-reducing anaerobic microorganisms have been isolated most frequently, and their diversity is comparable to that of sulfate-reducing microorganisms. The present chapter gives an overview of prokaryotes that reduce sulfate or elemental

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CHAPTER 1.22

Nitrogen compounds phosphate

Cell components

Organic

SO42– S0

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CO2

Free H S 2 Energy

Fig. 1. Metabolic principle of sulfate-reducing bacteria. As in other anaerobic bacteria, the main part of the electron donor is oxidized for energy conservation, and only a minor fraction is assimilated into cell mass. Catabolism (energy metabolism) is shown in black; anabolism (cell synthesis) is shown in red.

H2S

sulfur in their energy metabolism (see Fig. 1). Growth by reduction of other sulfur species is also included. Such bacteria have also been summarized as sulfidogenic bacteria (sulfideforming) bacteria (Zeikus, 1983; Lupton et al., 1984); however, strictly speaking, this term would also apply to putrefying bacteria that liberate sulfide from sulfur-containing organic molecules during their degradation. Sulfate- or sulfur-reducing microorganisms are long-established functional groups, like denitrifying, sulfur-oxidizing, methylotrophic or phototrophic bacteria. They are not necessarily coherent from the viewpoint of modern molecular systematics such as grouping based on 16S rRNA sequences. Nevertheless, the treatment of such functional groups besides molecular systematic groups is still the most appropriate basis for an understanding and comparison of physiological, bioenergetic and enzymatic properties and the roles of microorganisms in their natural habitat. Hence, the present chapter is mostly organized according to functional aspects, but it will distinguish between the phylogenetic domains and treat bacterial and archaeal sulfate-reducers and sulfur reducers separately.

Historical Overview Sulfate-Reducing Bacteria Meyer (1864) and Cohn (1867) first recognized the production of striking concentrations of H2S in aquatic habitats as a biologically mediated reduction of sulfate. Hoppe-Seyler (1886) demonstrated a complete oxidation of cellulose in anaerobic enrichment cultures with mud if gypsum (CaSO4) was provided as a source of sulfate; the latter was reduced to sulfide. Beijerinck’s (1895) investigations into microbial sulfide pro-

duction resulted in the first isolation of a sulfatereducing bacterium (named Spirillum desulfuricans), which was recognized as a strict anaerobe. The culture was isolated with malate and aspartate. Van Delden (1903a, 1903b) grew sulfatereducing bacteria on lactate, which is often still used for cultivation. The first thermophilic sulfate reducer with an optimal growth temperature of 55∞C was described by Elion (1925). Rubentschik (1928) observed a utilization of acetate and butyrate by sulfate reducers. In a comprehensive nutritional study, Baars (1930) demonstrated that Vibrio desulfuricans oxidized lactate or ethanol to acetate. Another type, Vibrio rubentschikii, was used in addition to acetate, propionate, butyrate and other compounds that were completely oxidized to CO2; unfortunately, this species was not preserved. Vibrio desulfuricans was the former Spirillum which finally became Desulfovibrio (Kluyver and van Niel, 1936; Stephenson and Strickland, 1931) observed an oxidation of H2 by sulfate reducers. The first described spore-forming sulfate-reducing bacteria were thermophiles named Clostridium nigrificans (Werkman and Weaver, 1927) and Sporovibrio desulfuricans (Starkey, 1938); they were later recognized as the same species (Campbell et al., 1957). In the 1950s and 1960s, principal insights into the biochemistry of sulfate-reducing bacteria were achieved. Desulfovibrio was the first anaerobe in which a cytochrome was detected (Ishimoto et al., 1954b; Postgate, 1953). Earlier, this type of pigment was thought to be associated only with O2 respiration. The type of cytochrome discovered in Desulfovibrio was termed c3. Investigations into the biochemistry of dissimilatory sulfate-reduction revealed differences from the pathway of assimilatory sulfate reduction known at that time (Lipmann, 1958). In Desulfovibrio, adenosine-5¢-phosphosulfate (APS) was not further phosphorylated to 3¢phosphoadenosine-5¢-phosphosulfate (PAPS), as

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

in the assimilatory pathway, but rather directly reduced to sulfite and AMP (Peck, 1959; 1962; 1965). Furthermore, electron transfer was demonstrated to be coupled to phosphorylation (Peck, 1966). A green protein, desulfoviridin, was first described by Postgate (1956) and subsequently recognized as sulfite-reductase. The mechanism of sulfite-reduction to sulfide was less understood. In addition to the electron acceptor sulfate, the metabolic fate of selected organic substrates, such as pyruvate and cysteine, was studied in sulfate-reducing bacteria (Senez, 1954a; Senez and Leroux-Gilleron, 1954b). Cultures of sulfate-reducing bacteria existing at that time oxidized their substrates (such as lactate, ethanol or malate) incompletely to acetate. Sulfate reducers formerly grown on acetate or higher fatty acids (Rubentschik, 1928; Baars, 1930) had not been preserved. In the 1960s, also a need for a proper classification of existing strains emerged. All sporeforming strains were classified or reclassified in the new genus Desulfotomaculum (Campbell and Postgate, 1965); the nonsporeforming, vibrio-shaped isolates were described as Desulfovibrio species (Postgate and Campbell, 1966). Later, nutritionally similar new mesophilic and thermophilic rod-shaped sulfate reducers were included in the genus Desulfovibrio (Rozanova and Khudyakova, 1974; Rozanova and Nazina, 1976); later, these sulfate reducers were reclassified as Desulfomicrobium and Thermodesulfobacterium, respectively. In the 1970s major advances were achieved in the characterization of various electron carriers, e.g., the resolution of the crystal structure of cytochrome c3 from Desulfovibrio (DerVartanian and LeGall, 1974). Furthermore, first evidence for a periplasmic location of hydrogenase emerged (Bell et al., 1974). In the field of biogeochemistry, new insights into the role of sulfate-reducing bacteria in natural habitats were rendered possible by the introduction of the radiotracer technique using 35 SO42- (Sorokin, 1972). More than 50% of the organic carbon in marine sediments was shown to be mineralized via sulfate reduction (Jørgensen and Fenchel, 1974; Jørgensen, 1977; Jørgensen, 1982). This process could not be explained by the incomplete substrate oxidation to acetate in the sulfate-reducing bacteria (Desulfovibrio and Desulfotomaculum species) known at that time. Anaerobic enrichment studies with various organic substrates lead to the recognition of diverse catabolic capacities including the degradation of aromatic organic acids in this group of microorganisms. Also, the capacity for acetate oxidation and complete mineralization of organic substrates, described in old reports

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(Hoppe-Seyler, 1886; Rubentschik, 1928; Baars, 1930), were confirmed to exist in sulfatereducing bacteria and found in several novel types of this group. Some new species were facultatively autotrophic. The diversity of the isolates required the establishment of a new Desulfotomaculum species (Widdel and Pfennig, 1977) and new genera, such as Desulfobacter, Desulfococcus, Desulfonema, Desulfobulbus and Desulfosarcina (Widdel and Pfennig, 1977; Widdel, 1980; Widdel and Pfennig, 1981b; Pfennig et al., 1981c; Widdel and Pfennig, 1982; Widdel et al., 1983). In the 1980s, main insights into enzymatic reactions and bioenergetics of entire metabolic pathways in sulfate-reducing bacteria were achieved and studies of functional genes began. Precise ATP balances of sulfate-reduction with H2 were calculated from chemostat studies (Badziong and Thauer, 1978; Nethe-Jaenchen and Thauer, 1984). In carbon metabolism, two alternative pathways for complete oxidation of acetyl-CoA, the citric acid cycle (Brandis-Heep et al., 1983; Gebhardt et al., 1983) and the oxidative CO-dehydrogenase pathway (Schauder et al., 1986; Schauder et al., 1989; Spormann and Thauer, 1988) were shown to be operative in distinct groups of sulfate-reducing bacteria that oxidized their substrate completely to CO2. In autotrophic sulfate-reducing bacteria (Widdel, 1980; Klemps et al., 1985; Brysch et al., 1987), the synthesis of acetyl-CoA from CO2 was demonstrated to occur via the reductive citric-acid cycle (Schauder et al., 1987) or the reductive COdehydrogenase pathway (Jansen et al., 1984; 1985; Schauder et al., 1989). Investigations into the metal clusters and cellular localization of hydrogenases led to the recognition of three different types of this enzyme in Desulfovibrio, the [Fe], [NiFe] and [NiFeSe] hydrogenase (Huynh et al., 1984a; Rieder et al., 1984; Teixeira et al., 1986; for summary see Fauque et al., 1988). First investigations into the molecular biology and genetics of sulfate-reducing bacteria included the study of plasmids (Postgate et al., 1984c; Postgate et al., 1986; Postgate et al., 1988; Powell et al., 1989) and genes for nitrogenase (Postgate et al., 1986; Kent et al., 1989), hydrogenase (Voordouw and Brenner, 1985a; Voordouw et al., 1985b), cytochromes (van Rooijen et al., 1989; Pollock et al., 1991), other redox proteins (Krey et al., 1988; Curley and Voordouw, 1988; Brumlik and Voordouw, 1989) and genes for biosynthetic enzymes (Li et al., 1986; Fons et al., 1987) in Desulfovibrio species. Also, genetic exchange systems were established for Desulfovibrio strains (Rapp and Wall, 1987; van den Berg et al., 1989; Powell et al., 1989). Furthermore, basic insights into the energy-mode of sulfate transport in various genera of sulfate-reducing bacteria were obtained

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(Cypionka, 1987; Cypionka, 1989; Warthmann and Cypionka, 1990). Attempts to enrich acetate-oxidizing anaerobes with sulfur-oxo anions other than sulfate led to the discovery of growth by disproportionation of sulfite and thiosulfate (Bak and Pfennig, 1987). The fact that anaerobic bacteria in natural habitats may be confronted with oxic conditions led to studies on the relation of various species of sulfate-reducing bacteria to O2 (Widdel, 1980; Cypionka et al., 1985; Dilling and Cypionka, 1990). Until the early 1980s, sulfate reducers were traditionally classified by phenotypic characteristics, such as nutrition, morphology and chemical, or biochemical markers, or both (Pfennig et al., 1981c; Postgate, 1984a; Widdel and Pfennig, 1984). Examples for applied chemotaxonomic markers are desulfoviridin (Postgate, 1959), lipid fatty acids (Boon et al., 1977; Ueki and Suto, 1979; Taylor and Parkes, 1983; Dowling et al., 1986), or menaquinones (Collins and Widdel, 1986). As the application of 16S rRNA sequence analysis became more and more common for the elucidation of natural relationships among microorganisms, this approach became decisive in the systematics of sulfate-reducing bacteria. The first comparative analysis of 16S rRNA sequence of a sulfate-reducing bacterium, Desulfovibrio desulfuricans, revealed relationships to Myxococcus and phototrophic purple bacteria (Oyaizu and Woese, 1985). A following comprehensive study based on the 16S rRNA oligonucleotide catalogs included the spore-forming Desulfotomaculum species and various nonsporeforming sulfate-reducing bacteria (Fowler et al., 1986). Desulfotomaculum was shown to branch with Gram-positive bacteria, as already indicated by the electron microscopy of the cell wall structure (Sleytr et al., 1969; Nazina and Pivavora, 1979). All other sul-

CHAPTER 1.22

fate reducers were found to affiliate with a branch of Gram-negative bacteria that also included the sulfur-reducing Desulfuromonas as well as Myxococcus and Bdellovibrio species. This branch of Gram-negative bacteria was termed the d-subdivision of the purple bacteria and their nonphototrophic relatives (Woese, 1987), even though a phototroph belonging to this subdivision has not been discovered thus far. Later, this phylogenetic assemblage became known as d-subclass of the Proteobacteria (Stackebrandt et al., 1988). Most described genera of sulfate-reducing bacteria affiliate with this subclass. Somewhat later, attempts were made to group the nutritionally diverse genera in meaningful higher taxa based on 16S rRNA sequences. First, two families were suggested within the sulfate-reducing bacteria of the dsubclass, the Desulfovibrionaceae and the Desulfobacteriaceae (Devereux et al., 1990; Widdel and Bak, 1992). However, the number of new isolates of sulfate-reducing and other bacteria and recognizable phylogenetic lineages within the d-subclass increased further. Today, a systematic structure of the d-subclass needs the establishment of several families and even orders. A novel thermophilic sulfate-reducing bacterium, Thermodesulfobacterium, was isolated in 1983 (Zeikus et al., 1983). Metabolically it resembled Desulfovibrio, however the lipids were ether-linked (Langworthy et al., 1983). Later, this organism was recognized as a deeply branching line of decent within the eubacteria, distant from the d-subclass of Proteobacteria (Henry et al., 1994). An earlier isolated thermophilic sulfate-reducing bacterium was recognized as a member of the same branch (Rozanova and Pivavora, 1988b). An overview of the major groups of sulfate-reducing bacteria and archaea (see following section) within the 16S rRNA-based tree of life is shown in Fig. 2.

Proteobacteria

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Microsporida

Fig. 2. Phylogenetic trees reflecting the relationships of groups of sulfatereducing bacteria to other organisms on the basis of 16S rRNA sequences. (A) Overview showing the three domains of life: (1), Eubacteria; (2), Archaebacteria; (3), Eukaryotes. The tree was adapted from AchenbachRichter et al. (1987) and Devereux et al. (1989). (B) More refined tree with genera. The tree was constructed using the ARB database and programs implemented therein (Ludwig et al., 1998). Scale bar represents 10 inferred nucleotide substitutions per 100 nucleotides.

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Sulfolobus solfataricus Archaeoglobus fulgidus Methanobacterium formicicum

Fig. 2. Continued

In the 1990s, many advances were achieved in the study of individual proteins, genes, degradative capacities and ecology of sulfate-reducing bacteria. Among the hydrogenases studied in various microorganisms, the first crystal structure was obtained from the enzyme in a sulfatereducing bacterium, Desulfovibrio gigas (Volbeda et al., 1995). Structural and functional investigations into hydrogenases continued steadily (Volbeda et al., 1996; Higuchi et al., 1997; Nicolet et al., 1999) and included the recognition of cyanide and CO as ligands of the active-site iron atom. Crystal structures of cytochrome c3 molecules from various Desulfovibrio sp. were also determined at high resolution (Matias et al., 1993; Czjzek et al., 1994; Fritz, 1999) and revealed a similar overall structure. Another contribution of general significance to biochemistry was the elucidation of the crystal structure of aldehyde oxidoreductase from Desulfovibrio gigas (Romão et al., 1995). The

crystal structure was not only the first to be resolved within the xanthine oxidase family, but also provided the correct structure of a widespread class of co-factors, the molybdopterins. The first crystal structure of a dissimilatory nitrate reductase was again determined from a sulfate-reducing bacterium, Desulfovibrio desulfuricans (Dias et al., 1999). Genetic studies were mainly carried out with Desulfovibrio desulfuricans and D. vulgaris because of the ease of cultivation and the applicability of antibiotics as selecting agents for mutants. Methods for the exchange of genetic material such as transduction, conjugation and transformation were further developed (for review see Voordouw and Wall, 1993b). Plasmids were constructed that can be applied as shuttle vectors for recombinant DNA (e.g., Rousset et al., 1998a). Transposons (Wall et al., 1996) and plasmids carrying the counterselectable marker sacB (Keon et al., 1997) were

664

R. Rabus, T.A. Hansen and F. Widdel

applied to sulfate-reducing bacteria to create mutants. In the 1990s, pure cultures of sulfate-reducing bacteria were isolated that could oxidize alkanes (Aeckersberg et al., 1991; Aeckersberg et al., 1998; So and Young, 1999a), toluene (Rabus et al., 1993; Beller et al., 1996), xylenes (Harms et al., 1999) or naphthalene (Galushko et al., 1999) completely to CO2. Furthermore, it was demonstrated that sulfate-reducing bacteria could grow with crude oil as the sole source of organic substrates (Rueter et al., 1994; Rabus et al., 1996), an aspect that contributes to our understanding of sulfide production in oil reservoirs and oil production plants. Anaerobic degradation of hydrocarbons as chemically sluggish molecules requires a suite of unusual reactions (e.g., the fumarate-dependent activation of toluene to benzylsuccinate; Beller and Spormann, 1997b; Rabus and Heider, 1998) as first discovered in denitrifiers (Biegert et al., 1996). In addition to hydrocarbons, other organic molecules were newly recognized as organic substrates for sulfate-reducing bacteria. Glycolate can be oxidized completely to CO2 by the novel sulfate reducer Desulfocystis glycolicus (Friedrich and Schink, 1995; Friedrich et al., 1996). Utilization of the sulfur compound dimethylsulfoniopropionate (DMSP) was demonstrated with several sulfate-reducing bacteria (van der Maarel et al., 1996a, b; Jansen and Hansen, 1998). Another type of novel sulfate-reducing bacterium was shown to oxidize a reduced inorganic phosphorous compound, phosphite (Schink and Friedrich, 2000). The introduction of molecular methods, especially those based on 16S rRNA sequences, into microbial ecology was also fruitful for the study of natural populations of sulfate-reducing bacteria. After the first construction of 16S rRNA-targeted probes for Desulfovibrio species (Amann et al., 1990) and other groups of sulfatereducing bacteria (Devereux et al., 1992), these and other probes were subsequently applied to biofilms (Ramsing et al., 1993; Santegoeds et al., 1999; Schramm et al., 1999), marine water columns (Ramsing et al., 1996; Teske et al., 1996), various sediments (Llobet-Brossa et al., 1998; Rooney-Varga et al., 1998; Sass et al., 1998; Sahm et al., 1999a), microbial mats (Fukui et al., 1999; Minz et al., 1999a) and an enrichment culture with crude oil (Rabus et al., 1996). Probe hybridization of rRNA after extraction or in whole cells, often in combination with counting series, confirmed the significance of sulfate-reducing bacteria in aquatic habitats, as shown in biogeochemical studies. Further approaches for the study of sulfate-reducing bacteria in habitats were based on reverse sample genome probing (Voordouw et al., 1991), hydrogenase genes

CHAPTER 1.22

(Wawer et al., 1997) or sulfite-reductase genes (Wagner et al., 1998; Minz et al., 1999b). Molecular methods in combination with cultivation and biogeochemical studies also provided basic insights into sulfate-reducing populations in cold sediments, which cover large areas of the ocean floor. Sulfate-reduction rates measured off Svalbard in the Arctic Ocean were comparable to those in marine sediments from temperate climate sites (Sagemann et al., 1998). Several previously unknown types of psychrophilic sulfate-reducing bacteria (e.g., Desulfotalea, Desulfofaba) could be isolated in pure cultures (Knoblauch et al., 1999a, b; Knoblauch and Jørgensen, 1999c) and shown to constitute a significant fraction of the natural cold-adapted population (Sahm et al., 1999b). The combination of pure-culture studies and molecular approaches also provided new insights into the ecology of gliding, filamentous sulfate-reducing bacteria, genus Desulfonema (Fukui et al., 1999).

Sulfate-Reducing Archaea When, during the early 1980s, several breakthroughs occurred in the discovery of novel, extremely thermophilic Archaea (see for instance Stetter, 1985), the novel isolates initially comprised methanogenic, fermentative, sulfurreducing and some aerobic microorganisms, but no sulfate reducers. Thermophilic sulfatereducing microorganisms known at that time were bacteria with temperature optima below 75∞C. In 1987, however, enrichment and isolation studies with samples from hydrothermal systems revealed the existence of archaeal sulfate reducers with a growth optimum of 83∞C (Stetter et al., 1987). The new sulfate reducer named Archaeoglobus fulgidus (Stetter, 1988) contains the cofactor F420, tetrahydromethanopterin (Stetter et al., 1987), and methanofuran (White, 1988; Gorris et al., 1991), which were known before only from methanogens. Furthermore, Archaeoglobus was shown to be phylogenetically more closely related to methanogens than to thermophilic archaeal sulfur-reducers or sulfur oxidizers (Achenbach-Richter et al., 1987). Further new species of the genus were A. profundus (Burggraf et al., 1990) and A. lithotrophicus (Stetter et al., 1993). Because the existing biochemical knowledge about mesophilic sulfate-reducing bacteria and methanogens could be applied to the study of Archaeoglobus, progress in the understanding of its metabolic pathways, enzymes and underlying genes was rapid. In the carbon metabolism, the pathway for complete oxidation of lactate to CO2 could be elucidated. It was recognized as an archaeal parallel of the CO dehydrogenase pathway in mesophilic sulfate-reducing bacteria

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

(Schauder et al., 1986; Schauder et al., 1989), with the involvement of the archaeal cofactors (Möller-Zinkhan et al., 1989; Möller-Zinkhan and Thauer, 1990; Schmitz et al., 1991; Klein et al., 1993; Schwörer et al., 1993). Also, enzymes in the transport of reducing equivalents were investigated (Kunow et al., 1994; Kunow et al., 1995). Autotrophic CO2 fixation in A. lithotrophicus was recognized to occur via the reductive CO dehydrogenase pathway (Vorholt et al., 1995), again a parallel to CO2 assimilation in sulfatereducing bacteria (Jansen et al., 1984; 1985; Schauder et al., 1989). The reduction of sulfate was shown to involve the same enzymatic steps as in bacterial sulfate reducers. Enzymes of the sulfate-reduction pathway in Archaeoglobus were purified and compared to the analogous bacterial enzymes, especially on the gene level (Speich and Trüper, 1988; Dahl et al., 1990; 1993; 1994; 1999a; Speich et al., 1994; Sperling et al., 1998; Sperling et al., 1999). In 1997, the complete genome sequence of A. fulgidus was published (Klenk et al., 1997). This was the first genome sequence of a sulfatereducing prokaryote.

Sulfur-Reducing Bacteria Biological reduction of sulfur to sulfide with endogenous or added organic electron donors has been reported several times since the end of the 19th century (Beijerinck, 1895; Starkey, 1937; Woolfolk, 1962; for overview see Roy and Trudinger, 1970). The reaction has been observed in bacteria, cell extracts, fungi, other plants, and in animal tissues. In several instances, the early observed processes of sulfur reduction appear to be by-reactions (incidental sulfur reduction) in an artificially created situation without bioenergetic or ecological significance. First evidence for sulfur reduction as the sole source of energy for microbial growth was furnished by Pelsh (1936) who enriched novel vibrioid bacteria from mud using sulfur and H2 as defined substrates. The first pure cultures definitely growing by sulfur reduction was Desulfuromonas acetoxidans, an obligately anaerobic mesophile using acetate as electron donor (Pfennig and Biebl, 1976). The bacterium was discovered as the chemotrophic partner in a deep-green phototrophic culture originally known as “Chloropseudomonas ethylica”; this culture was thought to be related to green sulfur bacteria, but differed from them by the ability to grow on acetate and even ethanol without addition of sulfide as electron donor. The actual process in this culture was elucidated as a sulfursulfide cycle involving a green phototrophic sulfur bacterium that oxidized sulfide to elemental sulfur, and Desulfuromonas that reduced sulfur

665

with organic compounds (Pfennig and Biebl, 1976). Desulfuromonas was also the first pure culture of an obligate anaerobe shown to oxidize acetate and other organic substrates completely to CO2 (Pfennig and Biebl, 1976); earlier, anaerobic acetate oxidation was only known in denitrifying bacteria. Subsequently, similar mesophilic bacteria including obligate sulfur reducers were isolated with organic compounds and sulfur (Pfennig, 1984; for overview see Widdel, 1988; for more recent classification see Finster et al., 1997b). Several of these sulfur reducers were shown to grow on acetate and fumarate. The formerly observed growth by sulfur respiration with H2 (Pelsh, 1936) was confirmed by isolation of a spirilloid bacterium (strain 5175) which in addition used formate (Wolfe and Pfennig, 1977). Fumarate was used as alternative electron acceptor. Subsequently, further morphologically similar, spirilloid bacteria with an anaerobic catabolism of fumarate (or aspartate) were recognized as facultative sulfur-reducing bacteria that oxidized H2 or formate. These were a tentative Campylobacter species (Laanbroek et al., 1977; Laanbroek et al., 1978), a spirillum isolated on lactate and DMSO (Zinder and Brock, 1978a), and Wolinella (formerly Vibrio) succinogenes (Macy et al., 1986). Neither Desulfuromonas nor the spirilloid sulfur-reducers were able to reduce sulfate. However, the capacity for growth by sulfur reduction was also detected in sulfate-reducing bacteria. Growth on lactate or ethanol in the presence of sulfur was observed with Desulfovibrio gigas (Biebl and Pfennig, 1977; Fauque et al., 1979), with an isolate tentatively named Desulfovibrio multispirans (He et al., 1986), and with nutritionally similar but rod-shaped, desulfoviridin-negative, sulfate reducers (Biebl and Pfennig, 1977) affiliating with the later proposed genus Desulfomicrobium (Rozanova and Nazina, 1976; Rozanova et al., 1988a). Later, anaerobes originally isolated as ferric iron-reducing bacteria were shown to be facultative sulfur reducers (Balashova, 1985; Myers and Nealson, 1988; Caccavo et al., 1994) and, vice versa, sulfur-reducing Desulfuromonas was shown to reduce ferric iron (Roden and Lovley, 1993). Furthermore, a Pelobacter species that had been originally isolated as a fermentative bacterium was recognized as facultative reducer of sulfur and ferric iron (Lovley et al., 1995c). A novel moderately thermophilic type of sulfur-reducing, acetate-oxidizing anaerobe was designated Desulfurella acetivorans (BonchOsmolovskaya et al., 1990). Furthermore, the thermophiles Aquifex (Huber et al., 1992), Ammonifex (Huber et al., 1996) and Desulfuro-

666

R. Rabus, T.A. Hansen and F. Widdel

bacterium (L’Haridon et al., 1998) were described as hydrogen-utilizing sulfur-reducing bacteria; Ammonifex was originally isolated as a nitrate-reducing bacterium. Natural relationships of sulfur-reducing bacteria were first investigated by 16S rRNA oligonucleotide cataloguing of Desulfuromonas (Fowler et al., 1986); it affiliates with the dsubclass of Proteobacteria and branches within completely oxidizing sulfate-reducing bacteria; the result was later confirmed by near-complete sequencing when similar species were classified as Desulfuromusa (Liesack and Finster, 1994). The phylogenetic branch that comprises spirilloid sulfur-reducing bacteria was termed the e-subclass of Proteobacteria. The first isolate (strain 5175; Wolfe and Pfennig, 1977) was classified as Sulfurospirillum deleyanum (Schumacher et al., 1992). Desulfurella species were recognized as a distinct branch within the e-subclass with no specific relationship to sulfatereducing bacteria or Desulfuromonas (Rainey et al., 1993; Miroshnichenko et al., 1998). The first biochemical studies of sulfurreducing bacteria were devoted to certain redox proteins and metal centers (Probst et al., 1977; Bache et al., 1983) as well as to the metabolism of acetate (Gebhardt et al., 1985). Acetate oxidation was shown to occur via the citric acid cycle, either with initial activation by CoA transfer from succinyl-CoA as in Desulfuromonas (Gebhardt et al., 1985), or with ATP-dependent acetate activation as in Desulfurella (Schmitz et al., 1990). For the investigation of the biochemistry and bioenergetics of sulfur respiration, Wolinella (formerly Vibrio) succinogenes was a highly suitable model organism. This bacterium had been originally isolated as a fumarate-respiring organism (Wolin et al., 1961). The experimental approaches and results from the detailed studies of the electron transport from formate (or H2) to fumarate in Wolinella as a model of anaerobic respiration (see e.g., Kröger and Winkler, 1981; Graf et al., 1985; Hedderich et al., 1999) provided an important basis also for investigations into sulfur respiration by this bacterium. Evidence was provided that polysulfide and not elemental sulfur is the actual electron acceptor (Klimmek et al., 1991; Schauder and Kröger, 1993; Schauder and Müller, 1993; Fauque et al., 1994), and there was increasing support for a periplasmic rather than a cytoplasmic orientation of the active site of polysulfide reductase, as in the case of formate dehydrogenase and hydrogenase in Wolinella (Schröder et al., 1988; Krafft et al., 1992; Schauder and Kröger, 1993; Krafft et al., 1995). The three subunits of the polysulfide reductase were analyzed with respect to bound cofactors (e.g., molybdopterin, FeS centers) and

CHAPTER 1.22

the underlying genes (Krafft et al., 1992; Krafft et al., 1995). A protein that increased the efficacy (viz. decreased the KM value) of polysulfide reduction was identified and termed Sud protein; it was suggested that Sud scavenges free polysulfide in the periplasm and transports it to the active site of the reductase (Kreis-Kleinschmidt et al., 1995; Klimmek et al., 1998).

Sulfur-Reducing Archaea In the early 1970s, the first extremely thermoacidophilic microorganisms were reported (Brock et al., 1972; Brierley and Brierley, 1982). The organisms classified as Sulfolobus were aerobic sulfur oxidizers. Somewhat later, they were recognized as members of a new “kingdom” of life termed “Archaebacteria” (Woese and Fox, 1977; Woese et al., 1978). These findings stimulated (in the early 1980s) the search for further, novel thermophiles under alternative conditions for enrichment cultures. Anoxic media were used that contained complex organic substrates, H2 as well as elemental sulfur, a potential electron acceptor known from mesophilic bacteria (see above). Indeed, novel extremely thermophilic archaea were detected that grew anaerobically and produced sulfide (Fischer et al., 1983; Stetter, 1982; 1983a; 1983b; Zillig et al., 1981; 1982; 1983), and the number of novel isolates increased steadily in subsequent years (for overview see e.g., Stetter et al., 1990; 1996). Several isolates seemed to reduce sulfur in a by-reaction or as mere electron sink to facilitate fermentation (Zillig et al., 1982; for more recent overview see Schönheit and Schäfer, 1995; Hedderich et al., 1999). Nevertheless, evidence for sulfur respiration as a mode of energy metabolism in archaea was clearly provided in cultures of Thermoproteus and Pyrodictium species that grew with H2 as the only electron donor in the absence of organic compounds (Fischer et al., 1983; Stetter et al., 1983b). Further, newly isolated archaea that definitely grow by sulfur respiration, viz on H2 and sulfur, were Stygioglobus azoricus (Segerer et al., 1991), Pyrobaculum islandicum (Huber et al., 1987) and Stetteria hydrogenophila (Jochimsen et al., 1997). A unique versatility in sulfur metabolism was found in new lithoautotrophic thermophilic isolates, Acidianus infernus (Segerer et al., 1985; Segerer et al., 1986) and Desulfurolobus (originally Sulfolobus) ambivalens (Zillig et al., 1985; Zillig et al., 1986) that grew aerobically by sulfur oxidation as well as anaerobically by sulfur reduction with H2. In carbon assimilation during sulfur reduction with H2, the reductive citric acid cycle and more recently the hydroxypropionate pathway were shown to be operative in Thermoproteus neutro-

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

philus (Schäfer et al., 1986) and Acidianus (Menendez et al., 1999), respectively. In the course of investigations into the sugar metabolism in several hyperthermophiles (for overview see Selig et al., 1997), pathways also were investigated in the sulfur-respiring, facultatively organotrophic Thermoproteus tenax (Siebers and Hensel, 1993). Evidence was provided for a non-phosphorylated Entner-Doudoroff pathway and a modified Embden-Meyerhof-pathway. Furthermore, complete oxidation of organic substrates via the citric acid cycle was demonstrated in the facultatively organotrophic sulfurrespiring species Thermoproteus tenax and Pyrobaculum islandicum (Selig and Schönheit, 1994). So far, these are the only extremely thermophilic sulfur-reducing microorganisms shown to couple sulfur reduction to complete mineralization of organic compounds, analogous to Desulfuromonas and Desulfurella (see above). The electron transport during sulfur reduction with H2 was studied in Pyrodictium brockii (Phil et al., 1992; Maier, 1996) and Pyrodictium abyssii (Dirmeier et al., 1998); these species employ different transport chains.

Overview of Principal Properties Sulfate-Reducing Bacteria and Archaea Sulfate-reducing bacteria gain energy for cell synthesis and growth by coupling the oxidation of organic compounds or molecular hydrogen (H2) to the reduction of sulfate (SO42-) to sulfide (H2S, HS-), as schematically shown in Fig. 1. Hence, sulfate-reducing bacteria are easily recognized by the production of high sulfide concentrations (with non-limiting electron donor and sulfate, usually in the range of several millimolar) concomitantly with growth and the strict dependence of this process on the presence of free sulfate. This process is also termed “dissimilatory sulfate reduction,” to allow clear differentiation from assimilatory sulfate reduction. Assimilatory sulfate reduction generates reduced sulfur for biosynthesis (e.g., of cysteine) and is a widespread biochemical capacity in prokaryotes and plants. Assimilatory sulfate reduction does not lead to the excretion of sulfide. Only upon decay (putrefaction) of the biomass is the assimilated reduced sulfur released as sulfide. The amounts of sulfide produced by dissimilatory sulfate reduction with a given amount of biomass is by orders of magnitude higher than the amount of sulfide liberated from the organic sulfur during putrefaction of the same amount of biomass. If the average formula of biomass is approximately written as that of a carbohydrate (CH2O), an amount 1,000 g

667

(33.3 mol) would yield 133 mol [H], and thus allow formation of 16.7 mol or 567 g H2S by sulfate reduction (8 [H] needed per SO42-). With the approximate natural content of 1% organic sulfur, the same amount of biomass would only yield 10 g of H2S if degraded by merely putrefying bacteria. The production of high concentrations of H2S often indicates the activity and presence of sulfate-reducing microorganisms in natural habitats. The presence of H2S is obvious by its characteristic smell, black precipitation of ferrous sulfide when iron minerals are present, and white patches of elemental sulfur as an oxidation product formed in contact with air. Such signs for the activity of sulfate reducers are often encountered if organic substances accumulate in the presence of sulfate under anoxic conditions. Growth conditions for sulfate-reducing microorganisms prevail in sediments of virtually all aquatic habitats, which may be cold, moderate or geothermally heated up to ca. 105∞C. But also flooded soils such as rice paddies and technical aqueous systems (as for instance sludge digestors, oil tanks or vats in the paper-making industry) may offer suitable growth conditions for sulfate-reducing microorganisms. From such habitats, in particular marine sediments, a great variety of sulfate-reducing microorganisms has been isolated. The classification of the major groups of sulfate-reducing microorganisms is today based on 16S rRNA sequence analysis. This method is usually relevant for the definition of the more refined taxa, viz. genera and sometimes species; nevertheless, phenotypic features such as nutritional capacities or chemotaxonomic properties may be decisive as well on the genus level, and in combination with DNA-DNA hybridization, in particular on the species level. Bacterial sulfate reducers fall into three major branches, the dsubclass of Proteobacteria with more than twenty-five genera, the Gram-positive bacteria with the genera Desulfotomaculum and Desulfosporosinus, and branches formed by Thermodesulfobacterium and Thermodesulfovibrio (Fig. 2). Sulfate reducers in the latter branch are thermophilic, whereas the two other branches comprise psychrophilic, mesophilic as well as thermophilic species. Currently recognized genera of sulfate-reducing bacteria and archaea are summarized in Table 1. Sulfate-reducing bacteria are morphologically diverse; cell forms include cocci, rods, curved (vibrioid) types, cell aggregates (sarcina-like) and multicellular gliding filaments. Sulfatereducing microorganisms are strict anaerobes, even though certain species may tolerate and reduce oxygen for a limited period of time. Many sulfate-reducing microorganisms can grow by

Sphere Oval (forms aggregates) Rod Multicellular filaments Vibrio Vibrio Straight or curved rod, sporulates

Desulfococcus Desulfosarcina

Desulfobotulus Desulfoarculus Desulfotomaculum

Desulfomonile Desulfonema

Oval or rod Oval Oval or vibrio Oval

Vibrio

Morphology

Desulfomicrobium Desulfobulbus Desulfobacter Desulfobacterium

Bacteriad Desulfovibrio

Genus

Optimum temperature (∞C) 34 35–39 30–38 50–65h

37 30–32

28–36 33

28–37 28–39 28–32 20–35

30–38

Desulfoviridina -

+ ±

+ -

-

+ + + ± ±

i i CAC CO CO CO c c i CO i or CO

SO32-, S2O32SO32-, S2O32S2O32-, 3-Cl-benzoate SO32-, S2O32SO32SO32-, S2O32S2O32- Fumarate

±

+ ±

+

+

Electron acceptors for growth (other than SO42-)

i

Oxidation of organic electron donorsb

SO32-, S2O32-, Fumarate SO32-, S2O32SO32-, S2O32-, NO3SO32-, S2O32S2O32-

H2

Acetate (+) ±

-f (+)

(+) (+)

+ (+)

-

Propionate (+) ±

ND +

+ +

+ (±)

-

Higher fatty acids + + ±

ND +

+ +

±

-

Ethanol +

-

+ +

± + ± ±

+

Lactate + ±

±

+ +

+ + -e ±

+

±

+

±

+ ±

±

Succinate, fumarate, and/or malate

Electron donorsc

±

-

-

-

±

Fructose, and/or glucose

Table 1. Morphological and physiological properties of the genera of sulfate-reducing bacteria and archaea.

±

+ ±

+ +

±

-

Phenyl-substituted organic acids

Methanol, alanine

3- or 4-Anisate -

Methanol, glycerol, glycine, alanine, choline, furfural Methanol, glutarate, glutamate, phenol, aniline, nicotinate, indole Acetone -

Others utilized by some species

668 R. Rabus, T.A. Hansen and F. Widdel CHAPTER 1.22

Vibrio Rod Oval Rod

Curved rod Rod

Rod

Thermodesulfovibrio Thermodesulfobacterium Thermodesulforhabdus Desulfacinum Desulforhopalus Desulforhabdus Desulfonatronovibrio Desulfonatronum Desulfohalobium Desulfofustis

Desulfocella Desulfocapsa Desulfobacca Desulfuromusa

Desulfospira Desulfobacula

Desulfofrigus

Morphology

Straight or curved rod, sporulates Vibrio Rod Rod Oval Oval Rod Vibrio Vibrio Rod Rod

Desulfosporosinus

Genus

Optimum temperature (∞C) 10

26–30 28

34 20–30 37 30

65 65–70 60 60 18–19 37 37 37–40 37–40 28

30–37

Desulfoviridina -

ND

ND

-

-

SO32-, S2O32Fe(III)-citrate

SO32-, S2O32S0, NO3- Fumarate DMSO Fe(III)-citrate SO32-, S2O32-, S0 ND

±

+ -

c c

c

ND

+ + + + + + + + +

+

i i c c

i i c ND i c ND i i i

SO32-, S2O32S2O32SO32SO32-, S2O32SO32-, S2O32SO32-, S2O32SO32-, S2O32SO32-, S2O32SO32-, S2O32-, S0 SO32-, S0

Electron acceptors for growth (other than SO42-)

i

Oxidation of organic electron donorsb

S2O32-

H2

Acetate +

+

+ +

+ + + +

-

Propionate -

-

+

+ + + +

-

Higher fatty acids +

+ -

+ ND ±

+ + ND ND ND ND ND +

ND

Ethanol +

+

+ -

+ + + + + + +

+

Lactate +

+ -

+

+ + + + + + + +

+

Succinate, fumarate, and/or malate +

+ +

+

+ + +

ND

Fructose, and/or glucose -

ND

-

ND +

-

-

± +

ND

ND ND ND ND +

+

Phenyl-substituted organic acids

Electron donorsc

(Continued)

Betaine, proline Toluene, p-cresol, benzaldehyde, benzoate, phenylacetate, p-hydroxybenzaldehyde, p-hydroxybenzoate -

Hexanol Glycolate, betaine, choline, triethanolamine, indole l-Alanine, 2-methylbutyrate -

3,4,5-Trimethoxybenzoate

Others utilized by some species

CHAPTER 1.22 Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes 669

Sphere

Rod Rod

Morphology

Optimum temperature (∞C) 82–83

7 10

Desulfoviridina -

-

Electron acceptors for growth (other than SO42-) SO32-, S2O32SO32-, S2O32Fe(III)-citrate

Oxidation of organic electron donorsb CO

i i +

+

H2

Acetate -g

-

Propionate ND

+ -

Higher fatty acids ND

-

Ethanol ND

+ + +

+ +

Lactate ND

+ +

+

-

ND

-

Starch, peptides

-

Others utilized by some species

b

Symbols: +, present; ±, present or absent; -, absent. Symbols: c, complete to CO2 via unknown pathway; CAC, complete oxidation via citric acid cycle; CO, complete oxidation via carbon monoxide dehydrogenase/C1 pathway; i, incomplete oxidation to acetate as an end product. c Symbols: +, utilized; (+), poorly utilized; ± , utilized or not utilized; (±) poorly of not utilized; -, not utilized; ND, not determined or not reported. d References: Desulfovibrio (Postgate, 1984b), Desulfomicrobium (Rozanova et al., 1988), Desulfobulbus (Widdel and Pfennig, 1982), Desulfobacter (Widdel and Pfennig, 1981b; Widdel, 1987), Desulfobacterium (Brysch et al., 1987), Desulfococcus (Widdel, 1980), Desulfosarcina (Widdel, 1980), Desulfomonile (DeWeerd et al., 1990), Desulfonema (Widdel et al., 1983), Desulfobotulus (Widdel, 1980), Desulfoarculus (Widdel, 1980), Desulfotomaculum (Widdel and Pfennig, 1977; Widdel and Pfennig, 1981b), Desulfosporosinus (Stackebrandt et al., 1997; Campbell and Postgate, 1965; Klemps et al., 1985), Thermodesulfovibrio (Henry et al., 1994), Thermodesulfobacterium (Zeikus et al., 1983), Archaeoglobus (Burggraf et al., 1990; Stetter et al., 1987; Stetter, 1988), Thermodesulforhabdus (Beederet al., 1995), Desulfacinum (Rees et al., 1995), Desulforhopalus (Isaksen and Teske, 1996), Desulforhabdus (Oude Elferink et al., 1995), Desulfonatronovibrio (Zhilina et al., 1997), Desulfonatronum (Pikuta et al., 1998), Desulfohalobium (Ollivier et al., 1991), Desulfofustis (Friedrich etal., 1996), Desulfocella (Brandt etal., 1999), Desulfocapsa (Janssen et al., 1996), Desulfobacca (Oude Elferink et al., 1999), Desulfuromusa (Liesack and Finster, 1994), Desulfospira (Finster etal., 1997a), Desulfobacula (Rabus et al., 1993), Desulfofrigus (Knoblauch et al., 1999b), Desulfofaba (Knoblauch et al., 1999b), and Desulfotalea (Knoblauch et al., 1999b). e Utilized by a few unnamed strains but not by the validly published species. f May be utilized with thiosulfate as electron acceptor. g For further description see Daumas et al., 1988; Min and Zinder, 1990; Nazina et al., 1988; and Widdel, 1988. h Thermophilic species.

a

Archaead Archaeoglobus

Desulfofaba Desulfotalea

Genus

Succinate, fumarate, and/or malate

Electron donorsc

Fructose, and/or glucose

Table 1. Continued

Phenyl-substituted organic acids

670 R. Rabus, T.A. Hansen and F. Widdel CHAPTER 1.22

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

utilizing sulfite or thiosulfate as alternative electron acceptors, which are also reduced to sulfide. Fewer species have been described to utilize elemental sulfur or nitrate as electron acceptors (for growth), which are reduced to sulfide or ammonia, respectively. The involvement of an external electron acceptor in the energy metabolism allows anaerobic growth even on highly reduced compounds that cannot be utilized by purely fermentative bacteria. Indeed, the electron donors of sulfate-reducing microorganisms include end products of fermentative bacteria. Bacterial sulfate reducers are known to utilize a great variety of low-molecular mass organic compounds, including mono- and dicarboxylic aliphatic acids, alcohols, polar aromatic compounds and even hydrocarbons. Growth with polymers, such as polysaccharides, as in the case of archaeal sulfate reducers, has not been observed. Oxidation of organic compounds may be incomplete, leading to acetate (often simultaneously with CO2) as an end product, or complete, leading entirely to CO2. In the case of lactate, a relatively common substrate, the two possibilities for its metabolism are as follows: 2-

2CH3 CHOHCOO- + SO4 Æ 2CH3 CHOO- + 2HCO3 + HS - + H + o DG ¢ = -160 kJ mol sulfate

(1)

2-

2CH3 CHOHCOO- + 3SO4 Æ 6HCO3 + 3HS - + H + DGo ¢ = -85 kJ mol sulfate

(2)

Incomplete oxidation of organic substrates is due to the lack of a mechanism for the terminal oxidation of acetyl-CoA. Because of this fundamental catabolic difference, it is common to distinguish between two physiological groups, the incomplete and complete oxidizers. However, these are purely physiological or functional groups that overlap only partly with molecular systematic groups. The energy gain from dissimilatory sulfatereduction is relatively low in comparison to aerobic respiration. For instance, the free energy change (DG∞) of the complete oxidation of acetate or lactate with sulfate as electron acceptor is -48 or -128 kJ, respectively, whereas acetate or lactate oxidation with O2 provides -844 or -1323 kJ, respectively (here caluclated per mol of the organic substrate). Accordingly, by far the greater part of the organic substrate (or of H2) consumed by sulfate-reducing bacteria is oxidized in the energy metabolism (Fig. 1), as is obvious from relatively low growth yields. Examples of measured dissimilatory growth yields (YSulfate, cell dry mass formed per mol sulfate

671

reduced) are as follows: Desulfovibrio vulgaris, H2 (with acetate and CO2 as carbon source), 8.3 g (Badziong and Thauer, 1978); Desulfobacter postgatei, acetate, 4.8 g (Widdel and Pfennig, 1981b); Desulfovibrio inopinatus, lactate (incompletely oxidized), 17.8 g (Reichenbecher and Schink, 1997); Desulfococcus multivorans, benzoate (completely oxidized), 6.2 g (Widdel, 1980); strain NaphS2, naphthalene (completely oxidized), 6.4 g (average; Galushko et al., 1999). The portions of the organic electron donors and carbon sources assimilated into cell material were ca. 9% (acetate) and 11% (lactate, benzoate, naphthalene). However, growth yields are not constants. They may be influenced by substrate limitation and resulting growth rate (Badziong and Thauer, 1978), the sulfide concentration (Widdel and Pfennig, 1977), and temperature (Isaksen and Jørgensen, 1996a; Sass et al., 1998b; Knoblauch and Jørgensen, 1999c). Variable growth yields of the same bacterial species (growing on one type of substrate) may be interpreted as a variable efficacy of coupling between electron transport and energy conservation, or a variable portion of the conserved energy (viz. ATP) that is needed for maintenance and hence does not contribute to net cell growth. As in other bacteria, there is no strict, causal connection between free energy changes and highest growth rates (mmax) that can be reached under optimum conditions. Still, the tendency has been often observed that electron donors which allow high free energy changes and involve simple, common enzyme mechanisms (e.g. H2, formate, ethanol, lactate, malate) allow, in principle, faster growth than electron donors that provide less energy and require more complicated, “unusual” enzyme mechanisms (e.g. aromatic compounds, alkanes). But there are exceptions. Some specialized species may utilize the former type of substrates (if used at all) more slowly than one of the latter. Growth rates observed with sulfate-reducing bacteria under optimal conditions (in synthetic media in the laboratory, with saturating or almost saturating substrate concentrations) cover a wide range, as illustrated with a few examples: Desulfovibrio vulgaris, H2, 0.15 h-1 (doubling time, 4.6 h; Badziong and Thauer, 1978); Desulfobacter species, acetate, 0.035–0.039 h-1 (doubling time 20–18 h; Widdel and Pfennig, 1981b; Widdel, 1987); strain NaphS1, naphthalene, ca. 0.004 h-1 (doubling time, 1 week; Galushko et al., 1999). The resulting highest specific sulfate reduction rates (Vmax = mmax/YSulfate) with H2, acetate and naphthalene were 18, 7.3–8.1, and 0.64 mmol sulfate per g cell dry mass and hour, respectively. Most sulfate-reducing bacteria tolerate more than 10 mM sulfide, as repeatedly shown during

672

R. Rabus, T.A. Hansen and F. Widdel

characterization of various species (for references see Table 1). Sulfate-reducing bacteria utilizing aromatic hydrocarbons formed as much as 20–25 mM sulfide before growth ceased (Harms et al., 1999; Rueter et al., 1994). In contrast, some Desulfotomaculum species appear to be more sensitive to sulfide, which affects their growth at concentrations of 4–7 mM (Klemps et al., 1985; Widdel and Pfennig, 1977). In comparison to bacterial sulfate reducers, archaeal sulfate reducers have been detected relatively recently and fewer species are known. As thermophilic microorganisms with optimal growth at temperatures around 80∞C or higher, archaeal sulfate reducers are less ubiquituous than their bacterial counterparts. Rather, archaeal sulfate reducers appear to be restricted to habitats like hydrothermal vents, hot springs and deep, warm oil reservoirs. So far, fewer substrates are known for archaeal than for bacterial sulfate reducers. However, archaeal sulfate reducers were shown to utilize the polymers, starch and peptides. Oxidation of organic compounds is always complete, in the case of lactate according to equation (2).

Sulfur-Reducing Bacteria and Archaea In addition to sulfate-reducing microorganisms, a variety of prokaryotes exists that reduce elemental sulfur (or other, lower oxidation states of this element) but not sulfate. Among the lower oxidation states, the element sulfur (often written as S0, S8) is probably the most widespread sulfur species in sediments and geological deposits. Many chemical and biological oxidation processes of H2S do not directly lead to sulfate (the highest oxidation state) but rather to elemental sulfur, which therefore may accumulate. Prokaryotes that reduce sulfur do not form phylogenetically coherent groups of bacteria or archaea. Many prokaryotes have been directly enriched and isolated with sulfur as an electron acceptor (e.g., Pfennig and Biebl, 1976; Wolfe and Pfennig, 1977; Bonch-Osmolovskaya et al., 1990; Stetter, 1985). Furthermore, the capacity for growth with sulfur as electron acceptor has been documented for bacteria that were originally isolated on the basis of growth with other electron acceptors such as manganese (IV) (Myers and Nealson, 1988) or iron (III) (Caccavo et al., 1994). Conversely, microorganisms isolated with sulfur are often able to reduce other electron acceptors such as nitrate, iron(III), or thiosulfate. In contrast to dissimilatory sulfate reduction, the capacity for sulfur reduction also has been observed in bacteria that grow definitely with O2 and which are,

CHAPTER 1.22

therefore, facultative anaerobes. However, many sulfur-reducing microorganisms are strictly anaerobic. Among the sulfate-reducing bacteria, only a few species can grow with elemental sulfur (Biebl and Pfennig, 1977; Table 1). Other sulfatereducing bacteria may produce some H2S in a byreaction not leading to growth when transferred from sulfate-grown cultures to media with crystalline (rhombic) or colloidal sulfur. Growth of many species of sulfate reducers is even inhibited by sulfur (e.g., Widdel and Pfennig, 1981b; Widdel et al., 1983; Bak and Widdel, 1986a; 1986b; Burggraf et al., 1990), probably because elemental sulfur as an oxidant shifts the potential of redox couples in the medium and cells to unfavorable, positive values. Analogous to capacities in sulfate-reducing bacteria, the oxidation of organic substrates in sulfur-reducing bacteria may be incomplete and lead to acetate as an end product (as for instance in Sulfos pirillum, Wolinella, Shewanella and Pseudomonas mendocina), or complete and lead to CO2 as the final product (as for instance in Desulfofuromonas or Desulfurella). Whereas bacterial sulfur reducers may be mesophilic or moderately thermophilic, archaeal sulfur reducers are all extremely thermophilic. Typical habitats of the hyperthermophilic sulfur reducers are solfataric fields, hot springs and hydrothermal systems in the deep sea, whereas mesophilic bacterial sulfur reducers can be isolated from almost every freshwater or marine sediment, or even from wet soil. Unlike sulfate reduction, the reduction of the lower oxidation states of sulfur is not always a respiratory process. The compounds may only serve as hydrogen sinks for a “facilitated fermentation,” or they may even be reduced in by-reactions without an obvious bioenergetic benefit. These processes vary, forming a spectrum ranging between true sulfur respiration and sulfur reduction as a mere by-reaction. A freshwater Beggiatoa was found to reduce stored sulfur under anoxic conditions with added acetate (Nelson and Castenholz, 1981). A certain increase in cell mass indicated that the process allowed a limited energy conservation. A Chromatium species and the cyanobacterium Oscillatoria limnetica was found to reduce photosynthetically formed intracellular or extracellular sulfur, respectively, in the dark under anaerobic conditions, using storage carbohydrate (van Gemerden, 1968; Oren and Shilo, 1979); growth did not occur. The reactions probably sustained a maintenance metabolism. However, it is not quite clear whether energy was gained only by substrate-level phosphorylation during sugar degradation or in addition by sulfur respiration.

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

We propose to apply the term “sulfur-reducing bacteria” to those bacteria in which sulfur reduction is associated with a respiratory type of energy conservation (sulfur respiration). An overview of the morphological and physiological properties of bacteria and archaea definitely capable of S0-respiration is provided in Table 2. Additional microorganisms that can reduce S0 to H2S, even though a respiratory function remains unclear, have been summarized by Hedderich et al. (1999).

Physiology, Biochemistry and Molecular Biology Sulfate-Reducing Bacteria Much of the research on sulfate-reducing microorganisms has been devoted to their unique metabolism in which five major aspects may be distinguished: 1) Sulfate reduction to sulfide, which is biochemically more complicated than O2 reduction in aerobic organisms, requires an array of enzymes. Like carbon and nitrogen, sulfur may occur in eight different oxidation states. In biochemistry, sulfur may form bonds to hydrogen, carbon and oxygen, but also chains with S-S-bonds. Oxidation states lower than +VI (sulfate) are rather reactive and may undergo interconversions or autoxidation even at room temperature. This reactivity complicates analyses of intermediates in sulfur metabolism, but also confronts research with interesting questions. 2) Sulfate-reducing bacteria utilize a wide variety of organic compounds. Even though these are of low molecular mass and relatively simple in their structure, their oxidation under anoxic conditions often involves biochemically intriguing reactions. 3) The flow of reducing equivalents ([H], electrons) from the electron donors to the electron acceptor is associated with the respiratory energy conservation, and a great variety of electron carriers seem to be involved. 4) Synthesis of cell material from most organic substrates is expected to proceed via pathways commonly known from other bacteria and therefore has not been a major field of research. However, the capacity of a number of sulfate-reducing bacteria for cell synthesis solely from CO2 (and mineral salts) during growth on H2 and SO42- as sole energy source has attracted particular attention. 5) A fifth main aspect, metabolic regulation, is widely unexplored in sulfate-reducing bacteria. In the study of all these aspects, molecular and genetic analyses are of increasing importance. Reduction of Sulfate to Sulfide Reduction of sulfate to sulfide is an eight-

673

electron step process that occurs via a number of intermediates. However, unlike many nitratereducing bacteria, sulfate-reducing bacteria usually do not excrete the intermediate oxidation states, but only the final product sulfide. Only in two cases, excretion by Desulfovibrio desulfuricans of minor concentrations of sulfite or thiosulfate has been reported (Vainshtein et al., 1980; Fitz and Cypionka, 1990); this does not necessarily indicate that thiosulfate is a direct intermediate. Sulfate Transport Because all enzymatic steps leading from sulfate to sulfide occur in the cytoplasm or in association with the inner side of the cytoplasmic membrane, sulfate has to be transported into the cell. Sulfate uptake in sulfatereducing bacteria is driven by an ion-gradient, as demonstrated in studies with Desulfovibrio species, Desulfobulbus propionicus and Desulfococcus multivorans (Cypionka, 1987; Cypionka, 1989; Cypionka, 1994; Cypionka, 1995; Warthmann and Cypionka, 1990). In the freshwater species (Desulfovibrio desulfuricans, Desulfobulbus propionicus), sulfate is transported simultaneously with protons, as revealed by instantaneous pH shifts in active cell suspensions upon addition of sulfate. In contrast, sulfate uptake in moderately salt-dependent species (Desulfovibrio salexigens, Desulfococcus multivorans) is driven by sodium ions (Warthmann and Cypionka, 1990; Stahlmann et al., 1991; Kreke and Cypionka, 1992). Cells grown at very limiting (e. g., micromolar) sulfate concentrations as in a chemostat (Cypionka and Pfennig, 1986) most likely transported sulfate with three protons or sodium ions, which allowed sulfate to accumulate by factors of 103 to 104 (Stahlmann et al., 1991). If the efflux of a neutral end product, H2S, is taken into account, sulfate transport is electrogenic under these conditions. The driving force for sulfate transport was mainly the electric component of the electrochemical potential and to a lesser extent the cation concentration gradient. There is evidence for an H+/Na+ antiporter which creates a sodium gradient across the cytoplasmic membrane of sulfatereducing bacteria (Varma et al., 1983; Kreke and Cypionka, 1992). With increasing sulfate concentration in the growth medium, the highaccumulating sulfate-transport system was no longer detectable. Instead, cells obviously produced a low-accumulating system causing sulfate concentration inside the cell by a factor not higher than 102, thus avoiding the buildup of deleterious sulfate concentrations. The latter system probably transported sulfate with two H+ or Na+ ions. At very high (28 mM) sulfate concentration as in seawater or most laboratory cultures, another regulation system seemed to attenuate

a

Wolinella succinogenes and similar spirilloid types Pseudomonas mendocina subsp. Geobacter sulfurreducens Pelobacter carbinolicus Sulfurospirillum arcachonense

Bacteria Desulfuromonas acetoxidans Desulfurella acetivorans Desulfovibrio gigas Desulfomicrobium species Dethiosulfovibrio peptidovorans Desulfitobacterium chlororespiransc Sulfurospirillum deleyianum

Species

30–37 35–36 35 35 26

Rod Rod Rod Curved rods

30 52–57 30–36 28–37 42 37 25–30

Optimum temperature (∞C)

Spirillum or vibrio

Rod Rod Vibrio Rod Vibrio Rod Curved spiral

Morphology

Oxidation of organic electron donorsa ND ND i i

i

c c i i i i ND

+ + +

+

NDb ND + +

H2

Acetate ND + +

-

+ + -

Lactate and/or pyruvate ND + +

+

± + + + +

Succinate, malate, glutamate Ethanol, butyrate, succinate 2,3-butandiol, acetoin,ethylene glycol glutarate, glutamate

Ethanol, propionate, succinate, glutamate Peptides, amino acids Butyrate Succinate, fumarate, malate aspartate, oxaloacetated Formate

Others utilized by some species

Sulfur + + + +

+

+ + + + + + +

Thiosulfate + -

+

+ + + + +

ND -

+

+ + + +

Sulfite

Electron acceptors

-

-

+ + -

Sulfate

Electron donors

ND + -

+

± + +

Fumarate

Table 2. Morphological and physiological properties of Bacteria and Archaea capable of respiratory reduction of elemental sulfur.

+ -

+

± +

Nitrate

+ +

+e

+

O2

674 R. Rabus, T.A. Hansen and F. Widdel CHAPTER 1.22

Lobed coccus Lobed coccus Long rod Disc with fibers Rod Lobed Coccus (irregular) Disk

Rod Rod Rod Rod

Morphology

Optimum temperature (∞C) 90 88 100 105 80 80 95 85

52–54 70 85 70

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

-f -f ND -f c -f -f -f

Oxidation of organic electron donorsa c -f ND

H2

Acetate -

+ -

Lactate and/or pyruvate ND -

ND +

Yeast extract

(With O2: sulfur) (With O2: sulfur) Yeast extract Yeast extract

(With O2: sulfur, thiosulfate) formate

Ethanol, stearate, palmitate

Others utilized by some species

Sulfur + + + + + + + +

+ + + ND

Thiosulfate + + ND +

+ ND -

Sulfite + + -

+ ND -

ND

ND +

ND ND ND ND ND ND -

ND ND -

ND -

+ +

Nitrate

+ + ND

+ -

O2

a

Symbols: c, complete oxidation (under anoxic conditions); i, incomplete oxidation; +, utilized; ±, utilized or not utilized; -, not utilized; ND, not determined or not reported. For further description and literature: Desulfuromonas acetoxidans, Desulfovibrio gigas and Desulfomicrobium species (Pfennig and Biebl 1976, 1981; Biebl and Pfennig, 1977; and Widdel, 1988; some data were personal communication from R. Bache and N. Pfennig); Desulfurella acetivorans (Bonch-Osmolovskaya etal., 1990; Schmitz et al., 1990); Dethiosulfovibrio peptidovorans (Magot et al., 1997); Desulfitobacterium chlororespirans (Sanford et al., 1996); Sulfurospirillum deleyianum (Wolfe and Pfennig, 1977; Schumacher et al., 1992); Wolinella succinogenes (Wolin et al., 1961; Macy et al., 1986); Pseudomonas mendocina subsp. (Balashova, 1985); Geobacter sulfurreducens (Caccavo et al., 1994); Pelobacter carbinolicus (Schink, 1984; Lovley et al., 1995); Sulfurospirillum arcachonense (Finster et al., 1997b; Stolz et al., 1999); Hippea maritima (Miroshnichenko et al., 1999); Desulfurobacterium thermolithotrophum (L’Haridon et al., 1998); Aquifex pyrophilus (Huber et al., 1992); Ammonifex degensii (Huber et al., 1996); Acidianus infernus (Segerer et al., 1985, 1986; Stetter et al., 1990); Sulfolobus ambivalens (Zillig et al., 1985, 1986 [the 1986 paper is about Desulfurolobus ambivalens]); Pyrobaculum islandicum (Huber et al., 1987; Selig and Schönheit, 1994); Pyrodictium occultum (Fischer et al., 1983; Stetter et al., 1983); Thermoproteus tenax (Zillig et al., 1981; Fischer et al., 1983; Schäfer et al., 1986; Stetter et al., 1990; Selig and Schönheit, 1994); Stygiolobus azoricus (Segerer et al., 1991); Stetteria hydrogenophila (Jochimsen et al., 1997); Thermodiscus maritimus (Fischer et al., 1993). b Not tested, but likely to be utilized. c Desulfitobacterium chlororespirans can also grow on lactate coupled to reductive dehalogenation of 3-chloro-4-hydroxybenzoate. d Utilized during fementative metabolism. e Low partial pressure (microaerobic conditions). f Obligate lithoautotrophs that do not oxidize organic compounds.

Hippea maritima Desulfurobacterium thermolithotrophum Aquifex pyrophilus Ammonifex degensii Archaeaa Acidianus infernus Sulfolobus ambivalens Pyrobaculum islandicum Pyrodictium occultum Thermoproteus tenax Stygiolobus azoricus Stetteria hydrogenophila Thermodiscus maritimus

Species

Sulfate

Electron acceptors

Fumarate

Electron donors

CHAPTER 1.22 Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes 675

676

R. Rabus, T.A. Hansen and F. Widdel

CHAPTER 1.22

even the low-accumulating system to prevent excess buildup of sulfate. Most likely, the sulfate transport systems operate near equilibrium (Cypionka, 1994; Cypionka, 1995). This means that the free energy from the gradient of the cotransported cations is not completely dispersed, but rather conserved more or less in the resulting sulfate gradient, rendering subsequent reactions of sulfate energetically more favorable than they would be at the lower ambient sulfate concentration. Hence, the consumption of 1/4–1/3 ATP equivalent per sulfate (assuming consumption of one electrogenically produced H+ ion, and a 3–4 H+/ATP stoichiometry of ATP synthase; Thauer and Morris, 1984; Stock et al., 1999) for sulfate transport at very low concentration must be regarded as energetically highly economic (Cypionka, 1995). The need for such reversible, energy-conserving transport processes in the catabolism is understandable in view of the relatively low ATP gain per mol sulfate. Sulfate uptake solely for biosynthesis (assimilatory sulfate reduction) differs completely from that in dissimilatory sulfate reduction. Sulfate transport in Escherichia coli for assimilation was shown to occur via an ABC transporter involving a periplasmic binding protein (Hryniewicz et al., 1990; Sirko et al., 1990). Such a mechanism for sulfate uptake is also likely in the cyanobacterium, Anacystis (Jeanjean and Broda, 1977). Sul-

Pyruvate

AeCoA + CO2

Fd(red) H2 (10Pa) H2 (10Pa)

fate uptake via ABC transporters for anabolic (assimilatory) purposes is irreversible (1 ATP/ SO42-); however, this dissipation of energy is negligible in view of the relatively low portion of reduced sulfur needed for cell synthesis (around 1% of dry mass). Activation of Sulfate The free sulfate dianion (SO42-) with its oxygen atoms in a tetrahedral arrangement is chemically sluggish and not easily reduced. The redox potential of the free anion pair SO42-/ SO32- is lower (E0¢ = –0.516 V) than redox potentials of most catabolic redox couples (Fig. 3). Before being reduced, sulfate is activated by ATP sulfurylase (Peck, 1959; Peck, 1962); the product is adenosine-5¢-phosphosulfate (APS), which is also termed adenylylsulfate. The ATP sulfurylase has been studied in several sulfate-reducing bacteria belonging to the genera Desulfovibrio and Desulfotomaculum (Fauque et al., 1991). Sulfate assimilation in nonsulfate-reducing bacteria and plants is also initiated by ATP sulfurylase; in the assimilatory pathways, APS either undergoes direct reduction, as in dissimilatory sulfate reduction, or phosphorylation to 3¢-phospho-adenosine-5¢phosphosulfate (PAPS) before reduction (Trudinger and Loughlin, 1981; Fischer, 1988; Peck and Lissolo, 1988).

E (V)

E (V)

–0.5

–0.5

–0.4

–0.4

Fd(ox) H+

2–

SO4

H+

NADPH

NADP+

–0.3

–0.3

Lactate

Pyruvate

–0.2

–0.2

Fdll(red)

Fdll(ox) –0.1

–0.1

SO3

–0.0

(2e) APS

MKH2 Succinate

–

SO3

–

MK

H2S

(6e)

Electrons Fumarate

–0.0

+0.1

+0.1

+0.2

+0.2

+0.3

+0.3

–

HSO+ 3 AMP

Fig. 3. Comparison of redox potentials of some important electron-donating and electron-accepting reactions in sulfatereducing bacteria. As mechanism of sulfite reduction to sulfide, a direct reduction with six electrons (6 e -) is assumed. For ferredoxin, an average of the E0¢ (–0.440 V) given by Fauque et al. (1991) and the E0¢ (–0.400 V) given by Thauer (1988) and Thauer et al. (1989b) is indicated. Abbreviations: Fd, ferredoxin; MK, menaquinone.

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

The equilibrium of the ATP sulfurylase reaction is far on the side of the reactants (Keq around 10-8; Akagi and Campbell, 1962), as has also been observed for the reaction in yeast (Robbins and Lipmann, 1958; Wilson and Bandurski, 1958). The hydrolysis of formed pyrophosphate (PPi) by a pyrophosphatase pulls the ATP sulfurylase reaction and thus favors APS formation (Wilson and Bandurski, 1958), according to the following reactions: 2-

SO4 + ATP + 2H + Æ APS + PPi DGo ¢ = +46 kJ mol

(3)

PPi + H 2 O Æ 2Pi DGo ¢ = -22 kJ mol

(4)

Sum reaction: 2SO4 + ATP + 2H + + H 2 O Æ APS + 2Pi (5) DGo ¢ = +24 kJ mol High pyrophosphatase activities were found in Desulfovibrio (Fauque et al., 1991), Desulfobulbus (Kremer and Hansen, 1988a), Desulfosporosinus orientis (Thebrath et al., 1989). Lower activities were observed in other Desulfotomaculum strains. However, earlier claims that PPi in this genus is used for an indirect phosphorylation of ADP via PPi:acetate kinase and acetate kinase (Liu and Peck, 1981a) have been questioned and are not supported by more recent experimental data (Thebrath et al., 1989). Still, use of PPi instead of ATP for certain phosphorylations during cell synthesis cannot be ruled out (Thauer, 1989a). Also, the possibility of energy conservation from PPi hydrolysis by using this reaction for proton translocation has been considered (Thebrath et al., 1989; Cypionka, 1995). On the other hand, any energy-conserving reaction that makes use of PPi has a certain reversible character and would diminish the pulling effect needed in reaction (4). Even with PPi hydrolysis, the thermodynamic equilibrium of the net reaction is still in favor of the reactants. With an assumed approximate concentration of sulfate, ATP and phosphate of a few millimolar (Thauer et al., 1977; Cypionka, 1995), the concentration of APS would have to be less than 0.1 mM to allow a net reaction according to equation (5). This indicates the need for effective scavenging of APS by reduction. One possibility to achieve this would be a close association of enzymes or enzyme complexes, in which molecules can be channelled between reaction centers and are not released into a cytoplasmic pool until the final product, sulfide, has been formed. However, such assumptions are presently speculative in view of experimental data.

677

Also in the activation of sulfate, the assimilatory and dissimilatory processes differ. Recent studies of assimilatory sulfate-reduction in E. coli K12 have revealed a novel mechanism for overcoming the unfavorable energetics of APS formation. In E. coli the intracellular concentration of PPi may be too high (ca. 0.5 mM; KukkoKalske et al., 1989) to allow formation of a substantial APS concentration. However, ATPsulfurylase in this organism was found to catalyze GTP hydrolysis in addition to APS formation. ATP-sulfurylase is a tetramer built of two heterodimers; each dimer consists of a CysN (53 kDa) subunit, which carries the GTPase activity, and a CysD (23 kDa) subunit, which carries the APS-synthesizing activity (Leyh et al., 1988; Liu et al., 1998). The presence of saturating concentrations of GTP stimulates APS formation by more than 100 fold (Leyh and Suo, 1992). The stoichiometry of GTP hydrolysis and APS formation was found to be 1 : 1 (Liu et al., 1998). The energy from GTP hydrolysis is transferred via conformational change to the formation of APS (Wei and Leyh, 1998; Wei and Leyh, 1999). The ATP-sulfurylase in E. coli, therefore, has been termed the “ATP sulfurylase-GTPase system”. The assimilatory ATP sulfurylase from E. coli and the dissimilatory enzyme from Desulfovibrio species also differ markedly on the structural level. A recent study on the composition of ATP-sulfurylase from two sulfatereducing bacteria, Desulfovibrio desulfuricans and Desulfovibrio gigas, demonstrated that here the ATP-sulfurylase is a homotrimer and contains the metals cobalt and zinc (Gavel et al., 1998). Reduction of APS APS is the actual electron acceptor, which is converted to sulfite or bisulfite and AMP. The E∞¢ of the APS/SO3- + AMP couple is –0.060 V. The actual redox potential may be more negative because of the expected low APS concentration (see above). APS reduction is catalyzed by a reductase that has been purified from Desulfovibrio strains (Bramlett and Peck, 1975; Lampreia et al., 1987), Desulfobulbus propionicus (Stille and Trüper, 1984), and Thermodesulfobacterium mobile (formerly Desulfovibrio thermophilus; Fauque et al., 1986). Presence of APS reductase was also demonstrated in Desulfobacter, Desulfococcus and Desulfosarcina (Stille and Trüper, 1984). Moreover, a type of this enzyme is found in some of the lithotrophic phototrophic purple and green bacteria and a few thiobacilli (Kelly, 1988; Fischer, 1988; Brune, 1989; Trüper, 1989). In these bacteria, APS reductase catalyzes the inverse reaction. All APS reductases are nonheme iron-sulfur flavoproteins. Purification of APS reductase from Desulfovibrio desulfuricans and Desulfovibrio

678

R. Rabus, T.A. Hansen and F. Widdel

CHAPTER 1.22

vulgaris under strictly anoxic conditions yielded highly active enzymes. The purified enzyme has a heterodimeric structure (ab), the total molecular mass being 95 kDa. Based on a characteristic motif in the primary structures, the a-subunit is proposed to carry one flavin adenine dinucleotide (FAD) molecule and the b-subunit to contain two [4Fe-4S] centers (Fritz, 1999). Two possible mechanisms for the reduction of APS to sulfite by APS reductase have been discussed. In the first proposed mechanism, the FAD group in the a-subunit is the active site. APS reacts with reduced flavin, FADH2, by a nucleophilic attack of the N5-atom. AMP is released and an FADH2-sulfite adduct is formed. Dissociation of sulfite with the binding electron pair then yields the oxidized FAD and protons (Peck and Bramlett, 1982a; Fig. 4). A formation of an FADH2-sulfite adduct as a possible intermediate during APS reduction was already suggested by Michaelis et al. (1970) and inferred from studies with artificial sulfite-flavin adducts

(Müller and Massey, 1969). More recent studies furnished increasing evidence for this mechanism. For instance, binding of APS, AMP and FADH2 could be demonstrated by spectroscopic measurements (Fritz, 1999). In the second proposed mechanism, a thiolate anion (R-S-) at the active site carries out a nucleophilic attack on the sulfur atom of APS. Such a reaction would result in the cleavage of the S-O-P bond, the release of AMP and the formation of a thiosulfonate group (R-SSO3-). Subsequent reduction of the thiosulfonate group with two electrons releases sulfite and restores the thiolate group of the active site (Fig. 5). The assumption that a thiolate group serves as active site was based on the finding that thiol-blocking agents inhibited the APS reductase from Desulfovibrio desulfuricans (Peck et al., 1965). However the recent experimental evidence that blockage of the thiol groups did not abolish but only reduce the activity rendered this mechanism unlikely (Fritz, 1999). In principle, the thiolate mechanism would resemble that in

NH2 N

N

O– O– 5¢ H2C O P O S O–

N

N

OH

OH

H

O

N 5

H3C

H+

NH 1 N H

N

H3C

O APS

O

O

O

O–

–

O

O

Adenosine O P O S O H 3C

N

H3C

N

O–

O NH

R FAD-reduced

O

N H

R NH2

2 [4Fe-4S]2+

N

N 2 [H]

O 5¢ H2C O P OH

N

N

OH

OH 2 [4Fe-4S]+ + 2H+

O

O O

AMP

–

O S O

O H3C

N

H3C

N

O NH

R FAD oxidized

N

O HSO3–

H+

H 3C

N

H3C

N

NH N H

O

R

FAD-sulfite adduct

Fig. 4. Proposed mechanism of APS reduction to sulfite with FAD as the catalytically active component. FAD is bound via a residue (R) to the enzyme. Electrons for the reduction of FAD are delivered by the [4Fe-4S] centers, the oxidation of which is indicated by the change of charge. Nucleophilic attack of N-5 results in binding of APS sulfur to reduced FAD. The FADsulfite adduct is formed upon release of AMP. Separation of sulfite from the enzyme yields oxidized FAD, which can reenter the reaction cycle. Abbreviations: APS, adenosine-5¢-phosphosulfate; FAD, flavin adenine dinucleotide.

CHAPTER 1.22 Fig. 5. Proposed mechanism of adenosine-5¢-phosphosulfate (APS) reduction to sulfite with a thiolate group as the catalytically active component. Nucleophilic attack of the enzyme-bound thiolate group leads to binding of APS. Upon release of AMP, a sulfite-enzyme adduct is formed. Reduction by two electrons (2 e-) allows separation of sulfite and regeneration of free thiolate.

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Adenosine

O

O

O P

O S

O–

O

O

Adenosine O P

O S

O–

S

679

Enzyme

–

O O–

O O–

O–

Adenosine O P –

O

O

Aps

AMP

O –

–

S Enzyme

O

SO32–

one of the assimilatory pathways. In the known assimilatory pathways, the sulfonate moiety from APS or PAPS is also transferred to a thiol, which can be glutathione or thioredoxin, to yield an organic thiosulfonate; this is either reduced to the corresponding organic persulfide (RSS-) or reductively cleaved with formation of sulfite, respectively (Trudinger and Loughlin, 1981; Imhoff, 1982; Fischer, 1988; Peck and Lissolo, 1988). In general, it is not known what electron donor is used in the cell to reduce APS. However, from Desulfovibrio vulgaris (strain Hildenborough), Chen et al. (1994d) isolated a flavin mononucleotide containing protein which not only catalyzed the oxidation of NADH by O2 with a concomitant formation of hydrogen peroxide (H2O2), but also fully reduced APS reductase with NADH as electron donor. Reduction of Sulfite Sulfite (:SO32-) or the protonated form bisulfite (tautomeric forms, [:SO2O-H]- and [H-SO2O:]-), which are approximately equally abundant at pH 7.0 (pKa2 = 6.99), are pyramidal molecules with free electron pairs at the sulfur and much more reactive than sulfate. Their metabolism needs no further activation by ATP. Early reports have suggested that bisulfite rather than sulfite is the actual substrate in the reduction to sulfide (Suh and Akagi, 1969; Drake and Akagi, 1977), and subsequently sulfite reductase has often been referred to as bisulfite reductase (Hatchikian, 1994). The reduction of sulfite (+IV) to sulfide (–II) by sulfite reductase involves the transfer of six electrons (equation 6). SO3

2-

+ 6e - + 8H + Æ H 2 S + 6H 2 O

(6)

The active centers of dissimilatory and assimilatory sulfite reductases (and nitrite reductases)

2e–

S Enzyme

S O

–

are characterized by two metallo-cofactors, a reduced porphyrin of the isobacteriochlorin class, the siroheme (Murphy and Siegel, 1973a; Murphy et al., 1973b; Murphy et al., 1974; Scott et al., 1978; Cole, 1988) and an iron-sulfur cluster ([FeS]). These metallo-cofactors function in the transfer of the electrons to the substrate, as indicated schematically in Fig. 6. Sirohemecontaining reductases have been isolated from a wide range of organisms. Siroheme was identified in assimilatory sulfite reductase from Escherichia coli (Murphy et al., 1973b), dissimilatory sulfite reductase from Desulfovibrio species (Murphy et al., 1973c), the dissimilatory “reverse” sulfite reductase of thiobacilli (Schedel et al., 1975; Trüper, 1994) and Chromatium (Schedel et al., 1979), and in the ammoniumproducing dissimilatory nitrite reductase from Escherichia coli (Jackson et al., 1981; Lin and Kuritzkes, 1987), higher plants (Hucklesby et al., 1976; Vega and Kamin, 1977), algae (Zumft, 1972) and fungi (Vega and Garret, 1975). Four major types of dissimilatory sulfite reductases are distinguished in sulfate-reducing bacteria, according to ultraviolet/visible absorption spectra and other molecular characteristics, the green protein desulfoviridin, the reddish brown colored desulforubidin and desulfofuscidin and P582 (Table 3; Fauque et al., 1991). Dissimilatory sulfite reductases generally have an a2b2 tetrameric subunit composition (Crane and Getzoff, 1996). However a third type of subunit (g) has been observed in a desulfoviridin-type of dissimilatory sulfite reductase in Desulfovibrio vulgaris (Pierik et al., 1992a) and Desulfovibrio desulfuricans strain Essex (Steuber et al., 1995), suggesting a hexameric structure (a2b2g2). The gsubunit is not encoded in the same operon as the a- and b-subunits and is not expressed coordinately with the a- and b-subunits (KarkhoffSchweizer et al., 1993). The molecular mass of

680

R. Rabus, T.A. Hansen and F. Widdel S

S

S

H2C

CHAPTER 1.22

Fe

Fe

CH2

FeS S

S Fe

S

H2C HO2C

CO2H S

HO2C

CO2H N

N Fe

H

N

CH3

N

CO2H HO2C CH3

H

CO2H HO2C

SO32–

Fig. 6. Prosthetic group of sulfite reductase. Two metallocofactors, the [FeS] cluster and siroheme, are covalently coupled via a sulfur bridge. Sulfite is ligated to the iron atom of siroheme from the opposite direction on the non-bridging side.

dissimilatory sulfite reductases ranges between 145 and 225 kDa. Desulfoviridin has been identified in virtually all Desulfovibrio species and has since been regarded as a taxonomic marker for this genus (Lee and Peck, 1971; Lee et al., 1973a; Postgate, 1984b). However, desulfoviridin also has been detected in Desulfococcus multivorans (Widdel, 1980) and most Desulfonema species (Fukui et al., 1999), which are unrelated to Desulfovibrio. Desulfoviridin is unique among the dissimilatory sulfite reductases in that it does not react with CO and contains siroheme (two per a2b2 holoenzyme) that is partly iron-free (viz. partly present as sirohydrochlorin). Siroheme and sirohydrochlorine are relatively easily released. The release of sirohydrochlorin is responsible for the red fluorescence in UV light of cells or extracts treated with dilute alkali (Postgate, 1956; Postgate, 1959). Siroheme prepared from desulfoviridin was found to catalyze the reduction of sulfite to sulfide and thiosulfate in the presence of artifical electron donors (Seki and Ishimoto, 1979). Analysis of the total iron content and spectroscopic investigations led to

architectural models of the siroheme and the [FeS] clusters in desulfoviridin. Hagen and coworkers reported that each molecule of desulfoviridin from Desulfovibrio vulgaris contains 20 iron ions and a demetallated siroheme. EPR and Mössbauer spectroscopy revealed an unusually high cluster spin of S = 9/2 of a putative [Fe6S6] prismane supercluster. Based on this finding, a superspin cluster was suggested with similarity to an [Fe6S6] prismane cluster observed in another redox protein from Desulfovibrio vulgaris (Marritt and Hagen, 1996). In the latter protein, four iron atoms probably form a core that is flanked on opposite sites by two iron atoms of more ionic character; the latter couple ferromagnetically through the core (Pierik et al., 1992b; Pierik et al., 1992c). Such a cluster should be able to accept more than one electron. However, other analyses of the crystal structure of the protein revealed the presence of only four Fe ions in a novel [4Fe-3S-2O] cluster structure (Arendsen et al., 1998). Furthermore, EPR spectra of dissimilatory sulfite reductase purified under strict exclusion of O2 yielded only weak signals, which also contradict the presence of a prismane-type super cluster (Fritz, 1999). Based on these findings, it is supposed that resonance signals previously thought to originate from a super cluster may actually result from oxidative damage of the [FeS] cluster of dissimilatory sulfite reductase. Desulfoviridin (containing 80% demetallated siroheme) from Desulfovibrio desulfuricans was reported to contain a total of 24 Fe ions (Steuber et al., 1995). Other reports on desulfoviridin from Desulfovibrio vulgaris furnished evidence for a total content of 10 Fe ions, and the presence of rhombic [Fe4S4] clusters (Moura et al., 1988; Wolfe et al., 1994). Desulforubidin was identified in a Desulfomicrobium strain (formerly regarded as a Desulfovibrio desulfuricans strain), which lacks desulfoviridin (Lee et al., 1973b), and in Desulfosarcina variablis (Arendsen et al., 1993). The Desulfomicrobium desulforubidin has been reported to possess an a2b2 structure (Moura et al., 1988; DerVartanian, 1994), whereas the corresponding enzyme from Desulfosarcina was demonstrated to have an a2b2g2 structure (Arendsen et al., 1993). Reports from the same authors on the total iron content and structure of the [FeS] cluster also suggest differences from the aforementioned results. Desulfofuscidin was purified and characterized from thermophilic sulfatereducing bacteria, Thermodesulfobacterium commune (Hatchikian and Zeikus, 1983; Hatchikian, 1994) and Thermodesulfobacterium mobile (Fauque et al., 1990). In both Thermodesulfobacterium species, the structure of desulfofuscidin was of the a2b2 type. In contrast to the two aforementioned dissimilatory sulfite reductases (des-

19(3)

[Fe4S4](2) [Fe6S6](3) +

SO32-(1,3)

S2-, S3O62-(1)

methylviologen(1)

(1)

10(4), 18(3)

[Fe4S4](4) [Fe6S6](5) -(1)

SO32-(1), NO2-(4) NH2OH(4) S2-(1), NH4+(4) S3O62-, S2O32-(6,7) methylviologen(1)

(1)

Acid-labile Sulfur

[FeS] clusters Reaction with CO Known substrate(s) Major products Minor products Electron donor (in vitro)

a

aSir, sulfite reductase.

(2)

(2)

Lee et al., 1973b Moura et al., 1988 (3) Arendsen et al., 1993

S3O62-, NH4+(1,2) S2-, S2O32-(1) methyl- or benzylviologen

15(3), 21(2)

10(4), 22(3)

Lee et al., 1973a Pierik et al., 1992a (3) Steuber et al., 1995 (4) Wolfe et al., 1994 (5) Pierik et al., 1992b,c (6) Akagi, 1983 (7) Lee and Peck, 1971

SO32-, NO2- NH2OH(2)

2(2)

2(4)

References

167(1) 190(2)

225(1)

Molecular weight (kDa) Number of sirohemes Total iron

Hatchikian and Zeikus, 1983

(1)

[Fe4S4] +(1)

16–17(1)

21(1), 32(2)

4(1,2)

a2b2(1)

a2b2a2b2 (%)(2,3)

628, 580, 408, 390, 279(1) a2b2 (%)(2,3) (n = 1–3) 226(1)

Absorption maxima (nm) Subunit structure

Desulfofuscidin Thermodesulfobactertium 693, 576, 389, 279(1)

Desulforubidin

Desulfovibrio Desulfosarcina 720, 580, 545, 392(1)

Desulfovibrio

Desulfoviridin

Organism

Properties

Table 3. Biochemical characteristics of sulfite reductases.

Trudinger, 1970 Akagi and Adams, 1973

(2)

(1)

SO32-, NO2NH2OH(1) S2-(1,2) S3O62-, S2O32-(2) methylviologen(1)

+(1)

54 matoms per g protein 15 matoms per g protein

145(1)

590, 545, 400(1)

700, 582, 392, 280(1)

Moura and Lino, 1994

(1)

S2methylviologen

SO32-

[Fe4S4](1) +

5(1)

5(1)

1(1)

27(2)

monomer

Desulfovibrio

alSiRa

Desulfotomaculum

P582

Type of sulfite reductase

Dahl et al., 1993 Dahl et al., 1994 (2)

(1)

methylviologen

Triper, 1994 Schedel et al., 1975 (3) Schedel et al., 1979 (2)

(1)

Siegel and Davis, 1974 (2) Siegel et al., 1982

(1)

SO32-, NO2NH2OH(1) S2-, NH4+ NADPH methylviologen SO32-, NO2-(1,2) SO32-

S2-(3) S2O32-, S2O3-(3) methyl- or benzylviologen

[Fe4S4]

4(1/b)

685

a2b2(1)

714, 587, 386, 280(1)

Escherichia

aSiR

[Fe4S4](1)

20(1), 47(3)

24(1), 51(3)

160(1) 280(3)

Thiobacillus Chromatium 700(3), 594, 393, 274(1) a2b2(1)a4b4 (3)

Reverse SiR

[Fe4S4]

20(2)

22–24(1)

2(1)

178.2(1)

593, 545, 394, 281(1) a2b2(1)

Archaeoglobus

Archaeal SiR

CHAPTER 1.22 Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes 681

682

R. Rabus, T.A. Hansen and F. Widdel

CHAPTER 1.22 -

H35 SO3 + [O3 S35 S-SO3 ]

A 6 e– SO32–

[O

3 H2O

3

H2S 8 H+

B 2 e– 3 H 2O 3 SO32–

2 e– S2O32–

S3O62– 6 H+

2 e–

SO32–

H2S 2 H+ SO32–

Fig. 7. Possible pathways of sulfite reduction to sulfide. (A) Direct reduction with six electrons without the formation of intermediates. (B) Trithionate pathway. The reduction occurs via three consecutive two-electron steps with the formation of tetrathionate and trithionate as intermediates.

ulfoviridin and desulforubidin), four instead of two siroheme cofactors per enzyme were found in desulfofuscidin. P582 was identified in the spore-forming Desulfotomaculum nigrificans (Trudinger, 1970; Akagi and Adams, 1973). Two different pathways for the reduction of sulfite to sulfide are discussed (Fig. 7): a sequential reduction in three two-electron steps with the formation trithionate and thiosulfate as intermediates, and a direct six-electron reduction without the formation of the aforementioned intermediates. Evidence for the first pathway, termed trithionate pathway (Fig. 7b) is mostly based on in vitro studies (Kobayashi et al., 1972; Akagi, 1983). In the in vitro experiments methylor benzylviologen were used as artificial electron donors; they were in a coupled system generated by reduction with hydrogen/hydrogenase. Under these conditions, trithionate and thiosulfate were identified in addition to sulfide as products of sulfite reduction. Under certain assay conditions, trithionate and thiosulfate were formed at concentrations similar to those of sulfide (Kobayashi et al., 1974). Also the enzymes in support of the proposed trithionate pathway, viz. trithionate reductase and thiosulfate reductase, were identified (Akagi et al., 1994). The purified desulfoviridin from Desulfovibrio gigas reduced sulfite with reduced methylviologen exclusively to trithionate (Lee and Peck, 1971). A “thiosulfateforming” enzyme was isolated from Desulfovibrio vulgaris which formed thiosulfate from bisulfite and trithionate. Labeling experiments with 35S demonstrated that the sulfur of formed thiosulfate originated from bisulfite and the inner S atom of trithionate, according to the following equation (Drake and Akagi, 1977).

35

S35 S

]

2-

-

2-

+ 2e - Æ

+ HSO3 + SO3

2-

(7)

Also purified from Desulfovibrio vulgaris was a “trithionate-reducing system”, which could form thiosulfate from trithionate and sulfite with flavodoxin (reduced by hydrogenase) serving as electron donor. In this system, a second protein was acting in close association with desulfoviridin and was required for trithionate formation (Kim and Akagi, 1985). A thiosulfate reductase that stoichiometrically reduced thiosulfate to sulfite and sulfide was purified from Desulfotomaculum nigrificans (Nakatsukasa and Akagi, 1969), Desulfovibrio gigas (Hatchikian, 1975) and D. vulgaris (Badziong and Thauer, 1980; Aketagawa et al., 1985). In summary, the stoichiometric formation of sulfite during the reduction of trithionate to thiosulfate and the reduction of thiosulfate to sulfite would add two loops to the pathway of sulfite reduction, as proposed by Kobayashi et al. (1974; Fig. 7b). Fitz and Cypionka (1990) reported the formation of trithionate and thiosulfate during reduction of sulfite with deenergized cells of Desulfovibrio desulfuricans. The occurrence of a trithionate pathway would be understandable from certain viewpoints of bioenergetics. The formation of trithionate would provide a relatively strong oxidant (E0¢ = +0.225 V) and thus a favorable acceptor even for high potential electron donors, as for instance from dehydrogenation of succinate (E0¢ = +0.030 V). The nature of the natural electron donor of the three two-electron reduction steps of the trithionate pathway has not been been resolved unequivocally (Peck and Lissolo, 1988). Furthermore, there are also arguments against a trithionate pathway (Chambers and Trudinger, 1975; Trudinger and Loughlin, 1981). The formation of trithionate and thiosulfate may be regarded as by-reactions. These may become dominant under in vitro conditions, for instance due to the relatively high concentrations of added bisulfite. Excess bisulfite or sulfite could react with intermediates bound to siroheme (Trudinger and Loughlin, 1981). Also, a reaction of bisulfite with formed H2S seems possible. Bisulfite and sulfide are known to react chemically to thiosulfate and thionates, especially at low pH (Heunisch, 1976). If the side-activities of certain proteins facilitated such a reaction under in vitro conditions, the produced sulfide would not accumulate but rather be scavenged to give rise to the observed oxo-anions. Trithionate and thiosulfate reductases may serve for utilization of their substrates from the environment or for scavenging them as byproducts of the bisulfite reductase reaction. Low concentrations (5–100 mM) of thiosulfate formed in deenergized

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

cells from added sulfite (Fitz and Cypionka, 1990) or in cells growing on sulfate (Vainshtein et al., 1980) also may be interpreted as byproducts resulting from a reversely operating thiosulfate reductase with sulfide and sulfite as reactants; the electrons from this low potential reaction (E0¢ = –0.402 V) could be easily consumed by other reductive processes. Evidence for the six-electron reduction of bisulfite to sulfide was achieved in a reconstitution assay with membrane-bound desulfoviridin, cytochrome c3 and hydrogenase, all from Desulfovibrio desulfuricans (Steuber et al., 1994); thiosulfate and trithionate were only detected in small amounts. This experiment also indicated that cytochrome c3 can act as electron donor for desulfoviridin, an observation that is topologically not yet understandable. The view of a six-electron reduction without the formation of free trithionate or thiosulfate as intermediates is favored if one compares sulfite reductases in dissimilatory and assimilatory sulfate metabolism and assumes that these enzymes, which are both siroheme proteins, employ in principle the same mechanism. In assimilatory sulfur metabolism, the assimilatory sulfite reductase generates sulfide for the synthesis of the sulfur-containing amino acid cysteine. Methionine and cofactors (like coenzyme A) derive their sulfur from cysteine. In contrast to dissimilatory sulfite reductase, none of the known assimilatory sulfite reductases (aSiRs) forms detectable amounts of trithionate or thiosulfate in vitro (Lee et al., 1973a; Peck and Lissolo, 1988). Thus aSiRs reduce sulfite with high fidelity directly via a six-electron reduction to sulfide. A sulfite reductase isolated from the sulfate-reducing bacterium Desulfovibrio vulgaris shared the high fidelity reduction of sulfite to sulfide with the aSiR from Escherichia coli. Therefore it was termed “assimilatory-like sulfite reductase” (Lee et al., 1973a). The enzyme has been studied in much detail. The aSiR from Desulfovibrio vulgaris was also functionally expressed in other Desulfovibrio hosts (Tan et al., 1994). The aSiRs from sulfate-reducing bacteria differ from the aSiR from Escherichia coli and the dissimilatory sulfite reductases from sulfate-reducing bacteria. The former are composed of only one polypeptide and do not form multimeric proteins; they have low-spin iron instead of high-spin iron and only one siroheme and [FeS]-cluster per molecule (Huynh et al., 1984a; Huynh et al., 1984b; Moura and Lino, 1994). Tan and Cowan (1991) proposed a mechanism for the the six-electron reduction catalyzed by aSiR, which may also serve as a working hypothesis to understand other sulfite reductases. The sulfur atom of sulfite binds the Fe2+ ion of the siroheme from the nonbridging face. A two-electron

683

reduction prepares the O-atom of the S-O bond for protonation so that a hydroxyl anion can be eliminated. Through repeated reduction by two electrons and subsequent protonation, the oxygen atoms are stepwise removed from the sulfur resulting in the formation of sulfide (Fig. 8). According to the model presented by Tan and Cowan (1991), the electrons for the reduction steps are “pushed” from the electron loaded [FeS]-clusters via the siroheme into the sulfite. In addition, the local environment in the sulfitebinding pocket may participate in the reduction reaction by providing protons from amino acid side chains to the O-atoms of sulfite. Such a mechanism would correspond to the “push and pull” paradigm, which has also been used to describe the cleavage of O-O bonds of peroxides by heme-containing oxygenases (Dawson, 1988; Poulos, 1988). Lui and Cowan (1994) have also proposed a six-electron reduction via a pushand-pull mechanism for dissimilatory sulfite reductase from Desulfovibrio vulgaris. In intact desulfoviridin, sulfite can only bind to reduced siroheme, whereas sulfite can bind to free siroheme in its oxidized state. These observations suggested a gating mechanism of dissimilatory sulfite reductase where a redox-linked structural transformation is required for substrate binding (Lui and Cowan, 1994). Insight into the mechanism of sulfite reduction and the structure of sulfite reductase have also benefited to a great extent from studies of the aSiR from Escherichia coli. This enzyme consists of eight flavoprotein subunits (a-subunits), which accept electrons from NADPH, and four hemoprotein subunits (b-subunits), which accept the electrons from the flavoprotein subunits and catalyze the six-electron reduction of sulfite to sulfide. Thus aSiR from E. coli has an overall a8b4-structure. Each hemoprotein-subunit carries one siroheme and one [Fe4S4]-cluster (Siegel et al., 1974; Siegel and Davis, 1974; Siegel et al., 1982). A chemical link between the siroheme and the [Fe4S4]-cluster was indicated by electronic exchange coupling observed by spectroscopic studies (Christner et al., 1984). The analysis of the crystallographic structure of the hemoprotein at a resolution of 3 Å suggested that a sulfur anion of a cysteine (Sg) covalently links the central iron in siroheme with one of the Fe ions in the cluster (McRee et al., 1986). A more recent analysis of the crystallographic structure of the hemoprotein at a resolution as high as 1.6 Å (Crane et al., 1995; Crane and Getzoff, 1996) demonstrated that the Sg is provided by Cys483. This bridge was found to be maintained in all reduction states of the enzyme studied so far on a structural level. The 1.6 Å structure also allowed recognition of further refined details of the structure. The hemoprotein consists of three

684

R. Rabus, T.A. Hansen and F. Widdel Fe2+ S

H+

L

Fe2+

Fe3+

S

S

Fe2+

Fe3+

SO32–

Fe2+ HO

CHAPTER 1.22

O–

S

OH–

HO

Fe2+ 2 e–

S Fe2+

O–

S

O

S

OH

O

H2S OH–

2 H+

Fe3+

Fe2+

S

S

Fe2+

Fe3+

Fe2+

S

S

Fe2+ S

2 e–

OH–

S OH

Fe3+ 2 e–

S Fe3+

H+

S O

Fig. 8. Suggested mechanism for the reduction of sulfite to sulfide by subsequent two-electron steps. The [4Fe-4S] cluster that is coupled to the Fe atom of siroheme via a sulfur bridge is represented by only one Fe-ion. The L represents the protein ligand that coordinates the Fe ion of siroheme. During catalysis, L is substituted by sulfite. Modified from Moura and Lino (1994).

domains that ligate the two metallo-cofactors at their interface. This interface is predominantly formed by b-sheets which are flanked at the outside by solvent-exposed a-helices. Domain 1 and 1¢ form a novel architecture reminiscent of a parachute and project harness hairpins into the interface-cofactor area. Domain 2 contributes the residues for the siroheme binding and positively charged residues that form the binding pocket for the anion substrate at the distal face of the siroheme. Domain 3 provides four cysteine residues (including the bridging ligand Cys483) to ligate the [Fe4S4]-cluster at the proximal side of the siroheme. The anion-binding pocket facing the distal side of the siroheme is remarkably rich in positively charged side chains, for instance Arg and Lys residues. Thus a strongly polarizing and proton-rich environment is established which may “pull” electrons of the S-O bond into the direction of the O-atom. Also water molecules could be positioned to interact directly with the anion substrate. Thus the structural details of the active site support the earlier model of a “push-and-pull” mechanism of the six-electron reduction of sulfite to sulfide. The structure of

the hemoprotein from E. coli is characterized by a vertical pseudo-twofold axis that relates an N-terminal sequence repeat (domain 1 and 2) to a C-terminal sequence repeat (domain 1¢ and 3); this suggests that the hemoprotein arose by gene duplication. Furthermore, analysis revealed the presence of five homologous regions in the sequence of the hemoprotein. Three of them (homology regions 1–3) encompass regions essential for the active center and for stabilization of the protein structure. Such homology regions have also been observed in dissimilatory sulfite reductases and therefore support the idea that dissimilatory sulfite reductases exhibit similar structure and also catalyze a six-electron reduction without formation of intermediates (Crane et al., 1995; Crane and Getzoff, 1996). There is a striking similarity between sulfite reductase and another enzyme with the capacity for a six-electron reduction, the ammonifying nitrite reductase (not to be confused with NOforming nitrite reductase in denitrifiers), which catalyzes the dissimilatory reduction of nitrite to ammonia (equation 8). Such types of nitrite

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

reductases also contain siroheme in the active center. -

+

NO2 + 6e - + 8H + Æ NH 4 + 2H 2 O

(8)

Cytochrome c nitrite reductase from Sulfurospirillum deleyianum not only catalyzes the sixelectron reduction of nitrite to ammonia, but also that of sulfite to sulfide. Interestingly, the analysis of the crystal structure of this enzyme (Einsle et al., 1999) revealed marked structural differences from the aSiR from E. coli. The former enzyme is a homodimer that is predominately composed of a-helices and contains 10 closely arranged hemes. Apparently, different structures and probably also varied mechanisms have evolved to accomplish a six-electron reduction. A membrane-bound cytochrome ccontaining nitrite reductase (also isolated from Desulfovibrio desulfuricans) catalyzes the sixelectron reduction of nitrite to ammonia as well as that of sulfite to sulfide (Liu et al., 1994; Pereira et al., 1996). Sequence analysis of the genes (dsr) encoding dissimilatory sulfite reductases from Desulfovibrio vulgaris, Archaeglobus fulgidus and Chromatium vinosum demonstrated that the three proteins are true homologues (Dahl et al., 1993; Hipp et al., 1997; Karkhoff-Schweizer et al., 1995). A more detailed study by Wagner et al. (1998) revealed that the evolutionary relationships derived from dsr sequences of sulfatereducing microorganisms were nearly identical to relationships inferred from the 16S rRNA sequences. The authors concluded that bacterial and archaeal dissimilatory sulfite reductases originated from a common ancestor. Dismutation of Sulfur Species A unique metabolic capacity of certain sulfate-reducing bacteria is growth by dismutation (disproportionation) of sulfite or thiosulfate, a process which may be formally described as an inorganic fermentation (Bak and Pfennig, 1987). The reactions are carried out by Desulfovibrio sulfodismutans, Desulfobacter curvatus and a so far unnamed species, strain NTA3, that grew significantly better by dismutation than by sulfate reduction. Growth by disproportionation of thiosulfate was also reported for an anaerobic bacterium, designated strain DCB-1 (Mohn and Tiedje, 1990a). Disproportionation of thiosulfate was demonstrated by radiotracer experiments in marine sediments and recognized as an important part of the sulfur cycle (Jørgensen, 1990; Jørgensen and Bak, 1991). 2-

2-

4SO3 + H + Æ 3SO4 + HS DGo ¢ = -58.9 kJ mol sulfite

(9)

2-

685

2-

S 2 O3 + H 2 O Æ SO4 + HS - + H + DGo ¢ = -21.9 kJ mol thiosulfate

(10)

A dismutation of elemental sulfur with standard activities of the products is thermodynamically unfavorable. However, because the activity of the insoluble, elemental sulfur is always equal to 1, the free energy of the reaction is strongly influenced by the concentrations of the products and the pH: 4S + 4H 2 O Æ SO4

2-

+ 3HS - + 5H +

(11)

Standard concentrations; pH = 7: DGo¢ = +10.2 kJ/mol sulfur SO42- and HS-, 0.001 M; pH = 7: DGo¢ = +6.9 kJ/mol sulfur SO42- and HS-, 0.001 M; pH = 8: DG = -11.3 kJ/mol sulfur A purely chemical dismutation of sulfur to H2S or polysulfide and oxygen-containing sulfur compounds was favored by elevated temperature and pH values above 7 (Belkin et al., 1985). If bacteria dismutated the formed sulfur-oxygen compounds to sulfide and sulfate, reactions (9) and (10) would result. Evidence for a microbial disproportionation of sulfur to sulfate and sulfide was provided by Thamdrup et al. (1993), who demonstrated this process in marine-enrichment cultures. The disproportionation of sulfur by these enrichment cultures was accompanied by a fractionation of the sulfur isotopes; sulfate was enriched in 34S and sulfide depleted in 34S (Canfield and Thamdrup, 1994). The sulfate reducer Desulfobulbus propionicus was the first microorganism shown to disproportionate sulfur in pure culture, even though growth under these conditions was very slow (Lovley and Phillips, 1994b; Fuseler and Cypionka, 1995). Two species of the new genus Desulfocapsa, D. thiozymogenes and D. sulfoexigens, grew well by disproportionation of sulfur. Both species required the presence of a sulfide scavenger (e.g., ferrihydrite) for growth with sulfur as sole source of energy and also can grow by disproportionation of thiosulfate and sulfite. Desulfocapsa thiozymogenes, but not Desulfocapsa sulfoexigens, can grow by reduction of sulfate to sulfide as the mode of energy conservation (Janssen et al., 1996; Finster et al., 1998). Evidence has been furnished that the disproportionation of sulfite or thiosulfate to sulfate and sulfide proceeds via a reversal of the reactions of dissimilatory sulfate reduction (Krämer and Cypionka, 1989). Thus, ATP sulfurylase and not ADP sulfurylase, which is found in many lithotrophic purple phototrophs (Brune, 1989; Fischer, 1988; ; 1989) and some thiobacilli (Kelly,

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1988; Kelly, 1989), is involved in the formation of sulfate from APS and allows “inorganic” substrate-level phosphorylation; it has not yet been established how stoichiometric amounts of PPi are formed for the conversion of APS to sulfate and ATP. Reducing equivalents are derived from conversion of bisulfite to APS, which has a relative positive potential (E0¢ = –0.060 V; see also Fig. 3). For the reductive part leading to H2S, shifting of these reducing equivalents by reversed electron transport seems to be necessary; this was indeed indicated by the sensitivity of the dismutation to uncouplers (Krämer and Cypionka, 1989). Electron Acceptors Other Than Sulfate Inorganic Sulfur Species Most sulfate-reducing bacteria can use thiosulfate and sulfite as electron acceptors in addition to sulfate (Table 1). Desulfotomaculum acetoxidans (Widdel and Pfennig, 1981b), Desulfonema magnum (Widdel et al., 1983), Desulfocella halophila (Brandt et al., 1999), and some sulfate reducers originally assigned to Desulfobacterium did not reduce sulfite in growth tests. Inasmuch as sulfite is assumed to be generally a free intermediate in dissimilatory sulfate reduction, the failure of sulfate-reducing bacteria to grow with sulfite at nontoxic concentrations may be due to the lack of a specific transport system. Oxoanions (other than sulfite and thiosulfate) have scarcely been tested in cultures of sulfate reducers. Desulfovibrio strains have been reported to reduce trithionate (S3O62-), tetrathionate (S4O62-), and dithionite (S2O42-) (Postgate, 1951; Ishimoto et al., 1954a; Fitz and Cypionka, 1990). Among the sulfate-reducing bacteria, some species such as of the genera Desulfohalobium, Desulfofustis, Desulfuromusa and Desulfospira can grow with elemental sulfur (see Sulfurreducing bacteria). Other sulfate reducers may produce some H2S in a by-reaction without growth after transfer of sulfate-grown cells to media with crystalline (rhombic) or colloidal sulfur. Growth of many species of sulfate reducers is even inhibited by sulfur (e.g.,Widdel and Pfennig, 1981b; Widdel et al., 1983; Bak and Widdel, 1986b), probably because sulfur as an oxidant shifts the potential of redox couples in the medium and cells into an unfavorable range. In sulfate-reducing bacteria able to grow with sulfur, its reduction is probably directly catalyzed by the tetraheme cytochrome c3 (Fauque et al., 1979; Fauque et al., 1980; Cammack et al., 1984). Sulfonates, DMSO Reduction of sulfonates by sulfate-reducing bacteria was first described by

CHAPTER 1.22

Lie et al. (1996). These authors demonstrated utilization of cysteate, isethionate (2-hydroxyethanesulfonate), and acetaldehyde-2-sulfonate by Desulfovibrio desulfuricans strain IC1. Isethionate was converted to sulfide and acetate. Cysteate was also used as an electron acceptor by strains of Desulfomicrobium baculatum DSM 1741 and Desulfobacterium autotrophicum. The former strain and Desulfovibrio desulfuricans ATCC 29577 also used isethionate. Desulfovibrio strain RZACYSA can use taurine (aminoethanesulfonate), cysteate, isethionate and aminoethanesulfonate as electron acceptors (Laue et al., 1997b). Cysteate and taurine also can be fermented by some sulfate-reducing bacteria (see below). Utilization of dimethylsulfoxide (DMSO) as an electron acceptor for growth of sulfatereducing bacteria resulting in the production of dimethylsulfide was first reported by Jonkers et al. (1996). Out of eight strains of sulfate reducers isolated from a marine or high-salt environment, five were shown to use DMSO; most of them were Desulfovibrio strains. In addition, one strain of the barophilic Desulfovibrio profundus was also shown to use DMSO by Bale et al. (1997); the same study also demonstrated DMSO reduction by the type strain of Desulfovibrio salexigens, which was reported by Jonkers et al. (1996) not to reduce this electron acceptor. Sulfate and DMSO were reduced simultaneously. Nitrate, Nitrite Nitrate is reduced by a few Desulfovibrio species (Seitz and Cypionka, 1986; Keith and Herbert, 1983; McCready et al., 1983; Mitchell et al., 1986), Desulfobulbus propionicus (Widdel and Pfennig, 1982) and Desulfobacterium catecholicum (Szewzyk and Pfennig, 1987; Moura et al., 1997). Nitrate may be preferred over sulfate (Seitz and Cypionka, 1986), or vice versa (Widdel and Pfennig, 1982). Dalsgaard and Bak (1994) showed that in an isolate from rice paddy soil, Desulfovibrio desulfuricans strain C4S, nitrate reduction was strongly inhibited by sulfide; at 0.46 mM sulfide, the specific growth rate was less than 10% of the maximum value, and no growth occurred at 0.75 mM sulfide. As the authors suggested, this implies that some negative results from growth tests of sulfate reducers with nitrate may be questioned because of the inclusion of 0.5 mM sulfide as a reducing agent in the media. In sulfate and sulfur reducers, the end product of nitrate reduction, which occurs via nitrite, is ammonia and not N2 as in denitrifying bacteria. The nitrate reductase of Desulfovibrio desulfuricans was purified and shown to be a monomeric

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

74-kDa protein with a [4Fe-4S] center and a molybopterin guanine dinucleotide cofactor (Moura et al., 1997). The crystal structure of this periplasmic enzyme has been determined (Dias et al., 1999); this is the first resolution of the three-dimensional structure of a nitrate reductase. Although bisulfite reductases also show activity toward nitrite, specific nitrite reductases appear to be involved in the subsequent reduction of formed nitrite to ammonium. A hexaheme cytochrome c3 acting as nitrite reductase and consisting of 62-kDa and 19-kDa subunits has been isolated from Desulfovibrio desulfuricans (Liu and Peck, 1981b; Moura et al., 1997). Iron (III) Whereas several non-sulfate-reducing members of the d-subclass of Proteobacteria, including sulfur-reducing bacteria, can grow with iron(III) compounds as electron acceptors, this capacity has only been occasionally observed in sulfate-reducing bacteria. Reduction of chelated iron(III) was demonstrated in enzymatic tests with several Desulfovibrio species, Desulfobacterium autotrophicum and Desulfobulbus propionicus (Lovley et al., 1993b), and in growth tests with Desulfovibrio profundus (Bale et al., 1997) and several psychrophilic species (Knoblauch et al., 1999b). Among the latter, Desulfotalea psychrophila also reduced insoluble (nonchelated) inorganic ferric iron (ferrihydrite); however, growth was not observed. Oxygen The study of the influence of O2 on bacteria with an anaerobic metabolism is an ecologically relevant and biochemically interesting topic. Exposure of anaerobic bacteria to O2 is a frequent, natural event in environments with fluctuating O2 penetration and at the anoxic/oxic interface. Furthermore, if oxic environments such as soils or oligotrophic sediments turn anoxic due to flooding or eutrophication, respectively, communities of anaerobic bacteria gradually become established; this is most likely to occur via passage of “inocula” through the oxic environment, as for instance oxic water. Studies of the effects of O2 on anaerobic bacteria include several aspects; these are, for instance, anaerobe tolerance of O2 and survival under oxic conditions, the possibility that O2 at low concentrations may even serve as electron acceptor and allow energy conservation, and the protection of cells against harmful effects. Pure cultures of sulfate-reducing bacteria in aerated media in laboratory experiments died off at different rates, depending on the species (Hardy and Hamilton, 1981; Cypionka et al.,

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1985; Abdollahi and Wimpenny, 1990). Simultaneous presence of sulfide sometimes increased the detrimental effect of O2 (Cypionka et al., 1985). This sensitivity to long-term oxic conditions suggest that permanently oxic waters or soil usually do not harbor nonsporeforming sulfate-reducing bacteria. Only endospores of Desulfotomaculum species may be present in such environments at significant numbers (Widdel, 1988). However, in dense aquatic microbial populations, nonsporeforming sulfatereducing bacteria were also observed in oxic zones. Studies on the natural distribution of sulfate-reducing bacteria revealed high numbers in zones that are exposed to rapid changes of the O2 concentration or that are even oxic over prolonged periods, e.g., in biofilms (Ramsing et al., 1993) and microbial mats (Canfield and Des Marais, 1991; Krekeler et al., 1997; Teske et al., 1998). The existence of anoxic microniches (Jørgensen, 1977) in such zones, which might explain the occurrence of active sulfate-reducing bacteria in oxic environments, is questionable; in microbial aggregates, the O2-reducing activity with the available electron donors was not sufficient to cope with O2 penetration (Plough et al., 1997). The effect of O2 on pure cultures of sulfatereducing bacteria in horizontal oxic/anoxic transition zones was studied in sulfidic agar with an organic electron donor under an oxic head space. In opposed O2-sulfide gradients, several species of sulfate-reducing bacteria exhibited growth, even though sulfate was absent (Widdel, 1980; Cypionka et al., 1985). However, there was evidence that the sulfate reducers used a chemical oxidation product of sulfide, most probably thiosulfate, as electron acceptor, without getting into direct contact with O2. The oxidation product was again reduced to sulfide. The resulting sulfur cycle mediated between the sulfate-reducing bacteria and the otherwise harmful O2 that served indirectly as final electron acceptor. Such a mediating cycle also may occur in natural habitats as long as electron donors are available. However, there is also evidence that sulfatereducing bacteria are able to utilize O2 directly. In experiments with cell suspensions of Desulfovibrio, O2 was shown to serve directly as electron acceptor for H2 oxidation, and to enable significant proton translocation (Dilling and Cypionka, 1990; Dannenberg et al., 1992). Growth due to O2 utilization has not been observed in these experiments. Nevertheless, owing to the high rates of O2 consumption, which were even higher than in aerobic bacteria (Krekeler et al., 1997; Kuhnigk et al., 1996), the respiratory activity of Desulfovibrio may be of

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considerable ecological relevance for the scavenging of O2 and an ATP gain for survival, if the habitat turns transiently oxic. The underlying mechanism of the buildup of a proton gradient with O2 as electron acceptor is not understood. It is true that cytochrome c3 can directly react with O2; however, as a periplasmic enzyme and electron acceptor of hydrogenase, cytochrome c3 reacting with O2 would not allow the generation of a proton gradient. Hence, O2 is expected to react with one or some of the redox proteins of the electron transport chain so that a proton gradient can be formed. From the viewpoint of thermodynamics, O2 is the most favorable of all electron acceptors and could replace any of the intermediates from the pathway of sulfate reduction. The problem lies in a controlled reaction of O2 that avoids instantaneous damage of proteins and redox centers by reactive oxygen species (e.g., superoxide or peroxide). In other experiments, O2 at concentrations as low as 0.24 to 0.48 mM were observed to support growth of Desulfovibrio vulgaris strain Hildenborough in lactate medium with a strongly limiting sulfate concentration (Johnson et al., 1997). However, it cannot be completely excluded that O2 was only indirectly reduced via a mediating sulfur cycle as suggested before (Cypionka et al., 1985). At concentrations above approximately 1 mM, O2 arrested growth of D. vulgaris (Johnson et al., 1997). In other experiments using four different strains of sulfate-reducing bacteria, the rate of sulfate reduction was strongly affected by an O2 concentration of 15 mM (Marshall et al., 1993). Reduction of O2 by sulfate-reducing bacteria may occur not only with electron acceptors directly utilized from the medium, but also with storage compounds. Desulfovibrio gigas and Desulfovibrio salexigens both can accumulate massive amounts of polyglucose during anaerobic growth with lactate and sulfate (Stams et al, 1983; van Niel et al., 1996; van Niel et al., 1998). Polyglucose utilization was shown to be involved in the survival under oxic conditions. In Desulfovibrio gigas, NADH produced during the breakdown of polyglucose was reoxidized by NADH:rubredoxin oxidoreductase, a dimeric flavoprotein consisting of a 27- and a 32-kDa subunit, and containing two molecules of each, FAD and FMN per enzyme molecule (LeGall and Xavier, 1996). The rubredoxin is oxidized with O2 at a flavoheme protein, yielding water as the end product (Gomes et al., 1997). A further argument for the assumption that O2 is not only a harmful agent, but at low concentrations, also a potential electron acceptor for respiratory energy conservation in sulfatereducing bacteria comes from the observation of aerotaxis in Desulfovibrio species. In medium

CHAPTER 1.22

with lactate and without (or limiting) sulfate in capillary tubes, Desulfovibrio species positioned themselves in bands at low O2 concentration (Johnson et al., 1997; Eschemann et al., 1999). Desulfovibrio oxyclinae formed ring-shaped bands around O2 bubbles (Krekeler et al., 1998). Band formation was dependent on the presence of an electron donor. Measurements of the O2 gradient with microelectrodes revealed that the side of the bands facing the bubbles was exposed to O2 concentrations of up to 50 mM, whereas the other side of the band was anoxic. This indicates an intensive O2 respiration within the band. Thus aerotactic band formation and O2 respiration can be regarded as a means to decrease the O2 concentration completely and restore anoxic conditions within a narrow zone (Eschemann et al., 1999). The attraction of sulfate-reducing bacteria by O2 at low concentrations is so far unique among anaerobic bacteria; they are usually assumed to be repelled by O2 (Armitage, 1997). A molecular key element of bacterial chemotactic response is the presence of methylaccepting chemotaxis proteins (MCPs), which receive and transmit the attracting or repelling signal. MCPs consist of a periplasmic N-terminal domain which binds the attractant or repellent, a transmembrane spanning segment, and a cytoplasmic C-terminal domain which functions as signal transducer. These proteins have been well studied in Escherichia coli and Salmonella typhimurium (Stock and Surette, 1996). A 73-kDa protein discovered in Desulfovibrio vulgaris (and named DcrA) shows in its C-terminal domain similarities to that of MCPs in E. coli; the Cterminal domain in the latter is the site of methylation. There is also evidence for a cytoplasmic location of the C-terminal domain of DcrA, in accordance with that of MCPs in E. coli (Dolla et al., 1992; Deckers and Voordouw, 1994b). In contrast, the N-terminal domain of DcrA did not exhibit significant sequence homology with known MCPs. The N-terminus of DcrA was found to harbor a c-type heme. Addition of O2 or the reducing agent dithionite resulted in a decrease or increase, respectively, in the methylation of DcrA. DcrA, a c-type cytochrome that was unknown before, may function in sensing O2 or the redox potential of the medium (Fu et al., 1994). To further elucidate the role of DcrA in chemotaxis, a knock-out mutant of the coding gene, dcrA, was constructed. However, phenotypic analysis of the mutant did not reveal a deficiency in aerotaxis (Fu and Voordouw, 1997). Subsequent analysis of a genome library of D. vulgaris strain Hildenborough revealed the presence of at least 11 additional dcr genes (dcrB to dcrL; Deckers and Voordouw, 1994a). Phylogenetic analysis suggested that the dcr family is

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

distinct from the mcp families in other eubacteria and arose early in evolution (Deckers and Voordouw, 1996). Also, proteins that might be involved in the detoxification of damaging oxygen species have been identified in sulfate-reducing bacteria. Superoxide dismutase and catalase activity have been detected in Desulfovibrio species (Bruschi et al., 1977; Hatchikian et al., 1977). Desulfoferredoxin (Moura et al., 1990) and neelaredoxin (Chen et al., 1994c) are mononuclear non-heme iron proteins that have been purified from Desulfovibrio species found to catalyze removal of superoxide (Romão et al., 1999; Silva et al., 1999b). Neelaredoxin is encoded in an operon with two additional open reading frames (ORFs) which putatively encode two chemotaxis proteins (Silva et al., 1999b). Interestingly, significant sequence similarities between desulfoferredoxin and neelaredoxin from Desulfovibrio and neelaredoxin and superoxide oxidoreductase from Pyrococcus furiosus were reported; the latter does not dismutate, but rather catalyzes a net reduction to H2O2 (Jenney et al., 1999). This observation indicates a mechanism of superoxide detoxification in sulfate-reducing bacteria that is different from the mechanism of superoxide dismutase. The genes rub and rbo, which code for rubredoxin and a putative rubredoxin oxidoreductase, respectively, were identified in Desulfovibrio vulgaris (Hildenborough) as one transcriptional unit (Brumlik et al., 1989). The rub-rob genes from Desulfoarculus baarsii complemented an Escherichia coli mutant that was deficient in superoxide dismutase (Pianzzola et al., 1996). The gene product Rob is suggested to scavenge superoxide not via dismutation as superoxide dismutase, but via a reductive mechanism using electron donors such as NAD(P)H (Liochev and Fridovich, 1997), possibly comparable to the abovementioned superoxide oxidoreductase. Definite aerobic growth of sulfate-reducing bacteria (viz. for an infinite number of generations in oxic media) has not been observed so far, despite their capacity to couple O2 reduction with energy conservation, their chemotaxis toward microaerobic zones, and their detoxification mechanisms. From the viewpoint of biochemistry, there is no obvious reason to assume that the capacity for dissimilatory sulfate reduction and aerobic growth in the same bacterium are mutually exclusive. Fumarate Some Desulfovibrio species ferment fumarate or malate. In the presence of an additional electron donor (e.g., H2 or formate), fumarate and malate are quantitatively reduced to succinate (Grossmann and Postgate, 1955; Miller

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and Wakerley, 1966; Barton et al., 1970; Wolfe and Pfennig, 1977), which represents a purely respiratory type of energy conservation (fumarate respiration; Graf et al., 1985; Kröger, 1987). Desulfovibrio desulfuricans reduced fumarate even prior to sulfate. Acrylate Reduction of acrylate as an alternative electron acceptor by sulfate-reducing bacteria was discovered by van der Maarel et al. (1996c). Acrylate can be formed in marine sediments by the cleavage of dimethylsulfoniopropionate (DMSP), an osmolyte of many marine algae; such a cleavage can be carried out by the acrylate-reducing sulfate reducer Desulfovibrio acrylicus. The DMSP lyase from this organism has been purified (van der Maarel et al., 1996b). Acrylate reduction to propionate also occurs in the presence of sulfate. Reductive Dehalogenation Reductive dehalogenation coupled to anaerobic bacterial growth was first demonstrated with 3-chlorobenzoate in a mixed culture (Dolfing and Tiedje, 1987). The organism responsible for the reaction was identified as a new type of sulfate-reducing bacterium named Desulfomonile tiedje (DeWeerd et al., 1990). 3-Chlorobenzoate and 3,5-dichlorobenzoate were used as electron acceptors for growth with formate as electron donor (Dolfing, 1990; Mohn and Tiedje, 1990b). Desulfovibrio strain TBP-1 can grow by coupling the oxidation of lactate to the reductive dehalogenation of 2,4,6tribromophenol to phenol (Boyle et al., 1999); other halogenated compounds that are used as alternative electron acceptors include 2-, 4-, 2,4and 2,6-bromophenol. Arsenate, Chromate and Uranium Anaerobic reduction of arsenate coupled to the oxidation of acetate was originally demonstrated with Chrysiogenes arsenatis. This strictly anaerobic bacterium cannot reduce sulfate (Macy et al., 1996). Desulfotomaculum auripigmentum is the first example of a sulfate-reducing bacterium that can grow with arsenate as a terminal electron acceptor (Newman et al., 1997b). This bacterium reduced arsenate to arsenite and preferred arsenate to sulfate when both were included in the medium; under such conditions, precipitation of As2S3 took place both intra- and extracellularly (Newman et al., 1997a). Two sulfatereducing bacteria, Desulfomicrobium strain Ben-RB and Desulfovibrio strain Ben-RA, can reduce sulfate and arsenate concomitantly (Macy et al., 2000). Studies on bacterial utilization of arsenate as electron acceptor have been summarized by Stolz and Oremland (1999b).

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Reduction of chromate(VI) with H2 as electron donor was observed with whole cells of Desulfovibrio vulgaris; reduction in cell-free extracts depended on cytochrome c3 (Lovley and Phillips, 1994a). Cytochrome c3 from D. vulgaris is also capable of uranium(VI) reduction (Lovley et al., 1993a). Electron Carriers and Possible Functions The reduction of one molecule sulfate to sulfide consumes eight electrons that are ultimately provided by the electron-donor substrate. Unlike the situation in aerobic respiration in mitochondria and bacteria, there is not one terminaloxidase analogue in sulfate reducers. These bacteria possess at least two simultaneously operating enzymes that are functionally analogous to a terminal oxidase, namely APS reductase and bisulfite reductase; two other enzymes, trithionate reductase and thiosulfate reductase, could have such a function in the case of stepwise sulfite reduction or with trithionate or thiosulfate as external electron acceptors. Unlike oxidases in aerobic respiration, the reductases of the sulfate-reducing bacteria were in most cases not found to be associated with the cytoplasmic membrane. In immunoelectron microscopy, the bisulfite reductases of Desulfovibrio vulgaris, D. gigas and Thermodesulfobacterium mobile (formerly D. thermophilus) and the APS reductases of D. vulgaris and D. gigas appeared to be cytoplasmic enzymes; only APS reductase of T. mobile was mainly membrane-associated (Kremer et al., 1988c). In the construction of electron flow models for chemiosmotic energy conservation by dissimilatory sulfate reduction, the frequent finding of bisulfite reductases and APS reductases in the cytoplasm and the possible involvement of two other reductases (trithionate reductase and thiosulfate reductase) are complicating factors. There is evidence that certain redox carriers have highly specific roles in the electron flow by transporting reducing equivalents to particular acceptors only (LeGall and Fauque, 1988; Peck and Lissolo, 1988; Fauque et al., 1991). Suggested mechanisms of energy conservation are discussed in connection with particular electron donors (see Energy Conservation in this Chapter). In the following subsections, a few characteristics of major redox proteins are presented. More detailed information is given by Fauque et al. (1990) and LeGall and Fauque (1988). Cytochromes Several different types of cytochromes, which differ in molecular mass, subunit composition and heme content, have been identified in sulfate-reducing bacteria (Widdel, 1988; Fauque et al., 1991; LeGall and Fauque, 1988).

CHAPTER 1.22

The physiological function of cytochromes with respect to their position in electron transfer is not yet completely understood. Principal types of cytochromes that have been recognized are the tetraheme cytochrome c3, the hexadecaheme high molecular mass cytochrome (Hmc), and the small cytochrome c553. The type of cytochrome that was named c3 has been identified in all Desulfovibrio species (Postgate, 1984a; LeGall and Fauque, 1988; Fauque et al., 1991), Desulfobulbus elongatus (Samain et al., 1986a) and both Thermodesulfobacterium species (Hatchikian et al., 1984; LeGall and Fauque, 1988; Fauque et al., 1991). Cytochrome c3 (Mr of ca. 13,000) consists of one polypeptide chain and contains four hemes with midpoint potentials ranging from –0.125 to –0.325 V; it is also termed tetraheme cytochrome c3. The ligands of each iron atom are two histidine molecules. The observed occurrence of cytochrome c3 in the periplasm (Badziong and Thauer, 1980; LeGall and Fauque, 1988; Fauque et al., 1991) has been confirmed by the signal sequence in the gene (Voordouw and Brenner, 1986). In cell-free systems, tetraheme cytochrome c3 is required for the reduction of ferredoxin, flavodoxin and rubredoxin by hydrogenase and apparently plays a key role in H2 metabolism (Fauque et al., 1991; LeGall and Fauque, 1988). Still, the mode of electron transfer by cytochrome c3 in vivo is unsatisfactorily understood. Cytochrome c3 may interact with the transmembrane spanning Hmc complex to channel electrons through the membrane into the cytoplasm (Voordouw, 1995). The crystal structures of cytochrome c3 from Desulfovibrio vulgaris Miyazaki (Higuchi et al., 1984), D. vulgaris Hildenborough (Matias et al., 1993), D. desulfuricans (Norway; Czjzek et al., 1994) and D. desulfuricans (Essex; Fritz, 1999) were resolved at resolutions lower than 2 Å. The three cytochromes had a similar overall structure with an extended a-helix and a short b-strand as the prominent secondary structure elements. The four heme groups are all solvent exposed and arranged in pairs (termed heme I/II and heme III/IV pairs). Conserved lysine residues surrounding heme IV are proposed to be essential for the contact between cytochrome c3 and the electron-delivering hydrogenase. D. africanus contains two different types of tetraheme cytochrome c3, one being acidic and another being basic. In contrast to the basic c3, the acidic c3 showed only poor reactivity towards either [Fe] or [NiFe] hydrogenase (Pieulle et al., 1996; Magro et al., 1997). An octaheme cytochrome c3 that is found in most Desulfovibiro species is structurally rather different from tetraheme cytochrome c3, but also can react with hydrogenase (Fauque et al., 1991;

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

LeGall and Fauque, 1988). Studies with mutants have indicated that heme IV is most likely the interactive heme in the cytochrome-hydrogenase complex, and that Tyr73 has an important structural function (Aubert et al., 1997; Aubert et al., 1998a). Octaheme cytochrome c3 may be involved in the supposed thiosulfate reduction of the trithionate pathway. A high molecular mass cytochrome c was isolated from Desulfovibrio vulgaris (Hildenborough); it had an estimated mass of 70 kDa and contained 16 heme groups (Higuchi et al., 1987). From the same organism, a dimeric 26-kDa cytochrome c3 (also referred to as cytochrome cc3) was isolated that possessed four identical heme groups in each subunit (Loutfi et al., 1989). A DNA probe designed from a partial amino acid sequence of cytochrome cc3 led to the identification of the hmc gene, coding for the hexadecaheme cytochrome Hmc in Desulfovibrio vulgaris (Hildenborough). In addition, the amino acid composition of the two cytochromes proved to be highly similar, thus suggesting that cytochrome c3 and Hmc are identical (Pollock et al., 1991). The hmc gene from D. vulgaris (Hildenborough) was overexpressed in D. desulfuricans G200 and the recombinant Hmc protein was purified. Studies on the arrangement of the heme-binding sites of this Hmc revealed that the protein contained three complete cytochrome c3-like and one incomplete c3-like domain, suggesting that Hmc arose via gene duplication (Bruschi et al., 1992). The hmc gene of D. vulgaris (Hildenborough) is part of an operon containing eight open reading frames, Orf1 to Orf6 (also termed hmcA to hmcF), Rrf1 and Rrf2. The open reading frame Orf1 represents the hmc gene. Based on sequence homologies, putative functions and cellular locations were suggested for the other open reading frames: Orf2 is a putative transmembrane protein containing four [FeS] clusters, Orf3 to Orf5 are membrane integral proteins, and Orf6 is a cytoplasmic protein containing two [FeS] clusters. It is proposed that Hmc and Orf2 to Orf6 are assembled in one transmembrane protein complex that functions in transferring electrons from the periplasm to the cytoplasm (Rossi et al., 1993). The two genes rrf1 and rrf2 code for regulatory proteins. Deletion of genes rrf1 and rrf2 resulted in an overexpression of the hmc operon and a more rapid growth on H2 and sulfate. From these results, it was concluded that the Hmc-complex mediates the electron transfer between periplasmic hydrogenase and the cytoplasmic enzymes involved in sulfate reduction (Keon et al., 1997). Even though Hmc from D. vulgaris (Hildenborough) can in principle accept electrons directly from [NiFe] hydrogenase, the rates of electron transfer are increased

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by the presence of cytochrome c3, suggesting that this cytochrome acts as a mediator between hydrogenase and Hmc (Pereira et al., 1998). However, an Hmc isolated from Desulfovibrio gigas could accept electrons directly from hydrogenase (Chen et al., 1994a). In the case of a transmembrane hexaheme cytochrome c from Desulfovibrio desulfuricans, a function of the protein as nitrite reductase could be demonstrated (Liu and Peck, 1981b). A “split-Soret” cytochrome, which is a dimer with two identical 26 kDa subunits and two heme groups per subunit, was isolated from Desulfovibrio desulfuricans (Liu et al., 1988). The complete amino acid sequence of this cytochrome c revealed that the C-terminal part contained the heme-binding site, similar to that in cytochrome c3, and an additional domain that could harbor a putative non-heme ironcontaining cluster (Devreese et al., 1997). During investigations on the natural electron acceptor of formate dehydrogenase of D. vulgaris, a small cytochrome with a mass of 6.5 kDa was isolated and (in accord with its absorption maximum) termed “cytochrome c553” (Yagi, 1969; Sebban et al., 1995). The purified protein could be reduced by formate dehydrogenase but not by hydrogenase (Yagi, 1979). Cytochrome c553 also can function as primary electron acceptor of lactate dehydrogenase (Ogata et al., 1981). Recognition of a leader sequence in the structural gene furnished evidence for a periplasmic location of cytochrome c553 (van Rooijen et al., 1989). Cytochrome c553 is a monoheme cytochrome with methionine and histidine as axial ligands (Fauque et al., 1991). The complete amino acid sequences of cytochromes c553 from D. vulgaris strains Hildenborough and Miyazaki revealed that the two proteins were not closely related (Nakano and Kikumoto, 1983). Apart from its small size, cytochrome c553 shows two further peculiar characteristics: 1) it has a low redox potential (ca. 0.01 V; Bertrand et al., 1982) and 2) it undergoes a conformational change during the transition from the oxidized to the reduced state (Senn et al., 1983). Structural analysis by means of NMR spectroscopy revealed that cytochrome c553 contains three conserved helices around the heme group, which resides in a cleft, and an additional fourth helix (Marion and Guerlesquin, 1992; Blackledge et al., 1995). In addition, the existence of two conformations of cytochrome c553 was recognized with NMR studies of the purified recombinant protein (Blanchard et al., 1993). The tyrosine residue Tyr64, which is positioned at the interface between the heme group and the central cleft of the protein, is thought to play a key role in structural stability (possibly affecting electron exchange with formate dehydrogenase;

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Blanchard et al., 1994; Sebban-Kreuzer et al., 1998a; Sebban-Kreuzer et al., 1998b). Further studies attempting to elucidate this electron transfer involve 15N labeling of cytochrome c553 and analysis with NMR techniques (Morelli et al., 1999). In addition to sulfate-reduction, Desulfomonile tiedjei DCB-1 can also employ reductive dehalogenation as a mode of energy conservation. A new type of cytochrome was found to be co-induced with the dehalogenating activity. This cytochrome is probably located toward the periplasmic aspect of the membrane because the protein was extracted from the membrane fraction and carries an N-terminal signal sequence. The coding gene of the new cytochromes was cloned by means of primers developed from the N-terminal sequence of the purified protein. Two c-type heme-binding motifs were identified in the C-terminus. However, the protein sequence was found to have no substantial similarities with sequences deposited in databases. Thus this protein is considered as a new c-type cytochrome (Louie et al., 1997). Ferredoxins Ferredoxins are very common in sulfate-reducing and sulfur-reducing bacteria (Probst et al., 1978; Bache et al., 1983; Gebhardt et al., 1983; LeGall and Fauque, 1988; Fauque et al., 1991). Several types have been described, but possible physiological roles are known only in a few cases. In Desulfovibrio gigas, ferredoxin I (E0¢ = –0.440 V), a protein with one [4Fe-4S] cluster, is active in the cleavage of pyruvate (viz. pyruvate:ferredoxin oxidoreductase reaction). In case of the assumed tetrathionate pathway, the midpoint potential of ferredoxin I would make it an appropiate electron donor for the thiosulfate reductase reaction. Ferredoxin II (E0¢ = –0.130 V) from D. gigas has one [3Fe-4S] cluster and has been suggested to function as electron donor in the reduction of bisulfite to sulfide (Fauque et al., 1991; LeGall and Fauque, 1988). There is evidence for an interconversion of the different clusters in these ferredoxins. In Desulfobacter as well as in the sulfur reducers (Desulfuromonas and Desulfurella), a ferredoxin is the acceptor in the 2-oxoglutarate dehydrogenase reaction (Gebhardt et al., 1983; Paulsen et al., 1986; Schmitz et al., 1990; Thauer, 1988; Thauer et al., 1989b). Flavodoxins In some but not all Desulfovibrio and Desulfomicrobium species, flavodoxins have been found. The two oxidation states, F/FH((E0¢ = –0.140 V) and FH(/FH2 (E0¢ = –0.440 V) have midpoint potentials comparable to those of ferredoxin I and II, and the corresponding proteins could replace each other in their function

CHAPTER 1.22

as electron carriers (Fauque et al., 1991). Flavodoxin was not active as electron donor for the purified thiosulfate reductase of Desulfovibrio vulgaris strain Miyazaki F (Aketagawa et al., 1985). The three-dimensional structure and the gene sequence of flavodoxin from Desulfovibrio vulgaris is known (Curley and Voordouw, 1988). Rubredoxins Rubredoxins are low molecular mass single-iron proteins (Mr ca. 6,000) which carry only electrons, like cytochromes and ferredoxins. They are present in all Desulfovibrio strains studied and also in Thermodesulfobacterium commune (LeGall and Fauque, 1988; Shimizu et al., 1989); the amino acid sequences of some of them have been determined, and the sequence of the gene in Desulfovibrio vulgaris (Hildenborough) coding for rubredoxin is known (Voordouw, 1988a; Voordouw, 1988b; Shimizu et al., 1989). Because of their rather positive midpoint potentials (–0.050 to +0.005 V), questions have been raised as to the physiological role of this protein in dissimilatory sulfate reduction (LeGall and Fauque, 1988; Brumlik and Voordouw, 1989). Kremer et al. (1988b) speculated about a role of rubredoxin as electron donor in the reduction of APS to bisulfite. However, such a role would be likely only if the actual potential of APS/HSO3- + AMP at the in vivo concentrations is more positive than the midpoint potential (–0.060 V), which may not be the case (see Reduction of APS in this Chapter). Experimental evidence for such a role has not been found so far. Rubredoxin may play a major role in channeling electrons to O2 consumption or O2 detoxification (See Oxygen in this Chapter). Rubrerythrin A high-potential redox protein, rubrerythrin (midpoint potential +0.23 V), has been purified from Desulfovibrio vulgaris and Desulfovibrio desulfuricans (Fauque et al., 1991); if this redox carrier is involved in sulfate reduction, a possible function that can be envisaged is in the reduction of trithionate to thiosulfate and bisulfite (E0¢ = +0.225 V). Recent studies indicate that the major role of rubrerythrin and nigerythrin may lie in protection against deleterious effects of O2. The proteins from Desulfovibrio vulgaris were shown to have NADH peroxidase activity (Coulter et al., 1999). Menaquinone All sulfate-reducing prokaryotes examined contain menaquinones (Collins and Widdel, 1986; Schmitz et al., 1990; Tindall et al., 1989). The number of isoprenoid units per side chain varies between 5 and 9. Terminal saturation of the side chain may occur in other bacteria

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

(Collins and Widdel, 1986). Judging from their general occurrence in sulfate-reducing microorganisms, menaquinones seem to be obligate components of electron transport chains. Involvement in the electron transport during oxidation of acetate (Kröger et al., 1988; MöllerZinkhan and Thauer, 1988; Möller-Zinkhan and Thauer, 1989; Thauer et al., 1989b; Figs. 6 and 8) or lactate (Fauque et al., 1991) has been discussed. Metabolism of Electron Donors and Energy Conservation A great variety of low molecular mass compounds serve a electron donors for dissimilatory sulfate reduction (and often simultaneously as carbon sources for cell synthesis). Many of these are products from the fermentative breakdown of biomass, which reflects the importance of sulfate-reducing bacteria as terminal degraders in anoxic, sulfate-rich habitats such as marine sediments. The study of several electron donors of sulfate-reducing bacteria is closely connected to investigations into structure and function of special enzymes such as hydrogenase. Furthermore, the study of electron donors has led to the discovery of previously unknown anaerobic pathways or capacities (e.g., a modified, anaerobic citric acid cycle, the oxidative C1/COdehydrogenase pathway, reactions at aromatic molecules or the capacity for alkane oxidation without O2). The bioenergetic processes of sulfate-reducing bacteria are determined by the electron donors. The catabolism of organic electron donors connected to the reduction of an external electron acceptor generally offers two advantages over a purely fermentative catabolism. First, with an external electron acceptor, substrate level phosphorylation can be performed to a larger extent than in purely fermentative metabolism in which a part of the organic substrate has to be sacrified for the disposal of surplus reducing equivalents (e.g., regeneration of NADP+). With external electron acceptors, substrate-level phosphorylation and growth is even possible with nonfermentable compounds, as for instance butyrate or higher fatty acids. In the vast number of sulfate-reducing bacteria that excrete acetate, substrate-level phosphorylation via phosphate acetyltransferase and acetate kinase can be expected. Substrate-level phosphorylation in Desulfobacter species occurs via ATP-citrate lyase, and in sulfate-reducing bacteria employing the C1/CO-dehydrogenase pathway most likely during formation of free formate from formyltetrahydropterin. Further substrate-level reactions may occur in the few species of sulfate-reducing

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microorganisms that grow with carbohydrates. Second and most importantly, the reduction of the external electron acceptor can be associated with an electron transport chain that allows generation of a transmembrane proton gradient and chemiosmotic ATP synthesis. In microorganisms utilizing organic compounds without an external electron acceptor, chemiosmotic energy conservation occurs only in special metabolic types, as for instance propionate-forming bacteria (Schink, 1988a) or methanogens. In sulfatereducing bacteria, numerous enzymes catalyzing redox-reactions as well as potentially electroncarrying proteins and menaquinones have been studied in detail, and electron transport chains have been proposed. However, there is no unifying theory of electron transport in sulfatereducing bacteria. In view of the various electron donors, metabolically diverse species and differences in the redox protein outfit, the development of a unifying model of electron transport is unlikely, except for steps in the pathway of sulfate reduction. In the following, physiological, enzymatic and energetic aspects of the utilization of various electron donors for sulfate reduction are presented. Molecular Hydrogen Molecular hydrogen (H2) is (besides acetate) a key intermediate in the natural mineralization of organic substances in sediments, sludge digestors and other anoxic ecosystems. Also the fact that many species of various genera of sulfate-reducing bacteria utilize H2 as sole electron donor (Table 1) reflects the ecological importance of the lightest of all molecules. Hydrogen at standard pressure is an energetically favorable electron donor (2 H+/H2, E0¢ = –0.414 V); the free energy change at various partial pressures is depicted in Fig. 11. Cell material during growth on H2 and sulfate may be synthesized from acetate and CO2 (chemolithoheterotrophic species) or alone from CO2 (see autotrophic species; Carbon Assimiliaton). Growth on H2 has been observed in many of the known genera of sulfate-reducing bacteria (Table 1). Hydrogenase activities have been demonstrated in strains of the genera Desulfovibrio (Fauque et al., 1991), Desulfobulbus (Samain et al., 1986b; Kremer and Hansen, 1988a), Desulfobacter, Desulfobacterium, Desulfosarcina (Schauder et al., 1986), Desulfotomaculum (Cypionka and Dilling 1986) and Thermodesulfobacterium (Fauque et al., 1992). Hydrogenase activity has even been found in Desulfobacter species that cannot grow on H2 (Lien and Torsvik, 1990); the role of the enzyme in such bacteria is unknown. Hydrogenases may

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CHAPTER 1.22

act not only in the uptake of H2 at various partial pressures (see last paragraph of this section), but also in the production of H2 during growth of certain species by fermentation or in syntrophic co-cultures (see Fermentative and Syntrophic Growth in the Absence of Sulfate in this Chapter). Hydrogenases catalyze the reversible heterolytic cleavage of H2 and oxidation of the resulting hydride ion, according to: H 2 ∫ H + + H - ∫ 2H + + 2e -

(12)

Detailed information on the biochemistry, coding genes and mechanism of function of hydrogenases (in sulfate-reducing bacteria) is so far only available from Desulfovibrio species. Hydrogenases are probably the most intensely studied enzymes in sulfate-reducing bacteria. Their investigation has significantly contributed to our understanding of hydrogenases in general. The first resolution of the three-dimensional structure of a hydrogenase was achieved with the enzyme from Desulfovibrio gigas (Volbeda et al., 1995). Based on their metal composition, three types of hydrogenases are distinguished in Desulfovibrio species, the [Fe] hydrogenases (Huynh et al., 1984a), the [NiFe] hydrogenases (Teixeira et al., 1986) and the [NiFeSe] hydrogenases (Rieder et al., 1984; Teixeira et al., 1987). All three types of hydrogenases have heterodimeric ab-structures and are mostly located in the periplasm (Odom and Peck, 1981a; Fauque et al., 1988). There are marked differences between the three types of enzymes (Table 4) with respect to their H2-uptake and H2-evolving activities, their sensitivity to CO, NO, NO2- and acetylene (e.g., He et al., 1989), and their molecular structures (Prickril et al., 1987; Fauque et al., 1988). The three types of hydrogenase are not uniformly distributed among Desulfovibrio species. Voordouw et al. (1990) analyzed the distribution of the hydrogenase encoding genes in 22 different Desulfovibrio species. The genes for the [NiFe] hydrogenase could be identified in all

tested strains, whereas the distribution of [Fe] and [NiFeSe] hydrogenases were limited. Individual strains may contain only one type of hydrogenase (e.g., [NiFe] hydrogenase in D. vulgaris strain Groningen), two types of hydrogenases (e.g., [NiFe] and [NiFeSe] hydrogenase in D. vulgaris strain Miyazaki; [NiFe] and [Fe] hydrogenases in D. desulfuricans strain El Agheila) or all three types of hydrogenases (in D. vulgaris strain Hildenborough). Genes coding for hydrogenases have been cloned and sequenced from various Desulfovibrio spp. and from Desulfomicrobium baculatum (Table 8). An extensive sequence comparison of hydrogenase genes including those from sulfate-reducing bacteria has been carried out by Wu and Mandrand (1993). The [NiFe] and [NiFeSe] hydrogenases from sulfatereducing bacteria were related to each other and also to [NiFe] hydrogenases from species from other subclasses of the Proteobacteria such as Rhodobacter, Rhizobium, Azotobacter, Escherichia or Wolinella. These hydrogenases were not related to [Fe] hydrogenases from Desulfovibrio species, which have their own line of enzymatic evolution. The [Fe] hydrogenases were purified from Desulfovibrio vulgaris strain Hildenborough (Huynh et al., 1984a), D. desulfuricans (Hatchikian et al., 1992) and D. fructosovorans (Casalot et al., 1998). In the case of [Fe] hydrogenases from D. vulgaris and D. desulfuricans, an atypical Fe-cluster and two ferredoxin-type [4Fe4S] clusters were identified. The atypical Fecluster, also known as the H-cluster, is assigned to the H2 activation site. The [4Fe-4S] clusters, which are also referred to as F-clusters, transfer electrons between the H-cluster and the external electron carrier (Adams, 1990). The crystal structure of the [Fe]hydrogenase from D. desulfuricans was the first to be determined of this type of hydrogenase (Nicolet et al., 1999). The threedimensional structure revealed that this hydrogenase displays a novel protein fold, and that the H-cluster is composed of a typical [4Fe-4S] clus-

Table 4. Brief overview of characteristics of different types of hydrogenases found in Desulfovibrio speciesa Catalytic activity H2 uptake H2 evolution Sensitivity to CO NO Nitrite Acetylene Molecular mass (kDa) a

Adapted from Fauque et al., 1991.

[Fe]hydrogenase

[NiFe]hydrogenase

[NiFeSe]hydrogenase

very high high

moderate moderate

low moderate

very high very high moderate no ~57

high high no high ~90

moderate very high moderate moderate ~81

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

ter bridged to a binuclear Fe center as the active site. The two Fe ions at the active site probably possess CO and CN- as binuclear ligands, as found in [NiFe] hydrogenases. The structural analysis of the [Fe] hydrogenase corroborates the earlier finding that one of the two active-site irons could be ligated by intrinsic CN- and CO (Pierik et al., 1998). In contrast to the [NiFe] hydrogenases, the binuclear active site as well as the [4Fe-4S] clusters in [Fe] hydrogenase reside on one subunit. Channel-like paths have been identified that allow the transport of protons and H2 to or from the active site buried in the center of the protein. The second subunit of the [Fe] hydrogenase from D. desulfuricans forms a belt around the other subunit. The [Fe] hydrogenase from the anaerobic Gram-positive bacterium C. pasteurianum has an active center similar to the one in the D. desulfuricans enzyme, even though the former consists of only a single polypeptide (Adams, 1990; Peters et al., 1998; Peters, 1999; Cammack, 1999). The hydA and hydB genes coding for the large and small subunits, respectively, of [Fe] hydrogenase in D. vulgaris (Hildenborough) and D. vulgaris subsp. oxamicus are highly homologous (Voordouw et al., 1989b); however, there is no significant homology between the [Fe] hydrogenases and the [NiFe] hydrogenases (see next paragraph). A gene probe for the [Fe] hydrogenase did not hybridize with the DNA of sulfate-reducing bacteria without a [Fe] hydrogenase (Voordouw et al., 1987). Deckers et al. (1990) demonstrated that D. vulgaris strain Miyazaki F lacks the [Fe] hydrogenase genes. In D. fructosovorans a new type of [Fe] hydrogenase, which reacts with NADP+, was identified; it may be regarded as a fourth type of hydrogenase present in Desulfovibrio. The NADP+-reducing hydrogenase is assumed to be a heterotetrameric enzyme complex that is encoded by the hndABCD genes (Malki et al., 1995). Mutants with deleted hndABCD genes showed reduced hydrogenase activity (Malki et al., 1997). Homology studies implicated that HndA and HndC form the NADP-reducing moiety, and that HndD harbors the H2activating site of a [Fe] hydrogenase; the function of HndB is presently unknown. The purified HndA subunit contains a [2Fe-2S] cluster which belongs to the family of [2Fe-2S] ferredoxins (DeLuca et al., 1998a). Studies with antisera raised against the four putative subunits overexpressed in (and purified from) Escherichia coli demonstrated that the active NADP+-reducing hydrogenase in the sulfate reducer is indeed a complex, even though the complex itself has not been purified so far (DeLuca et al., 1998b). Thus the NADP+reducing hydrogenase appears to differ

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structurally from the three other types of hydrogenases. The [NiFe] hydrogenases have been purified from D. desulfuricans (Krüger et al., 1982), D. gigas (Moura et al., 1982), D. multispirans (Czechowski et al., 1984) and D. africanus (Niviere et al., 1986). A [NiFe] hydrogenase was also isolated from the thermophilic sulfate reducer Thermodesulfobacterium mobile (Fauque et al., 1992). The [NiFe] hydrogenase from D. gigas has been studied most intensively. Analysis of the coding genes, hynA and hynB, suggested that the large subunit (62 kDa) carries the Ni ion and that the small subunit (26 kDa) could ligate at least two [FeS] clusters due to the presence of 12 cysteines (Voordouw et al., 1989a). Spectroscopic analysis of [NiFe] hydrogenase from D. gigas indicated the presence of two [4Fe-4S] clusters, one [3Fe-xS] cluster, one Ni ion and one unknown redox component, which was hypothesized to be a special Fe ion (Huynh et al., 1987; Albracht, 1994). The structure of the [NiFe] hydrogenase from D. gigas was determined at 2.85 and 2.54 Å resolution (Volbeda et al., 1995; Volbeda et al., 1996). The two subunits interact extensively, and the large subunit has a unique topology. The presence of Fe as the second metal ion, besides Ni, in the active site of the large subunit was demonstrated. The distance between the two metal ions was suggested to be around 3 Å. A coordination of intermediate species of H2 between the two metal ions (Ni and Fe) in the active site is suggested to function in catalysis (Volbeda et al., 1995). The Ni ion is anchored to the protein via sulfur bridges from two cysteine residues (Cys65 and Cys530) and coordinately bound to the Fe ion again by two sulfur bridges provided by Cys68 and Cys533. The Fe ion possesses three intrinsic dinuclear ligands, which were demonstrated to be two CN- groups and one CO molecule (Pierik et al., 1999). High resolution X-ray structural analysis (1.8 Å) of the [NiFe] hydrogenase from Desulfovibrio vulgaris (Miyazaki) indicated that this protein is similar to the [NiFe] hydrogenase from Desulfovibrio gigas with respect to the folding pattern, the arrangement of the metal center and the probable presence of SO, CO and CNas dinuclear ligands of the Ni-Fe center (Higuchi et al., 1997). These dinuclear ligands generate unusual infrared bands, which have been observed in several [NiFe] and [Fe] hydrogenases from Desulfovibrio species and other microorganisms like Chromatium vinosum (Bagley et al., 1995; van der Spek et al., 1996). Thus [NiFe] hydrogenases appear to contain NiFeCO(CN)2 as prosthetic group, the finding of which would be unprecedented in the study of biological systems (Fig. 9; Happe et al., 1997); the function of the dinuclear ligands remains unclear. The three

696

R. Rabus, T.A. Hansen and F. Widdel O N

N C

C

C Fe

Cys

S

S

Cys

Ni S Cys Fig. 9. Suggested prosthetic group of [NiFe] hydrogenases. Three dinuclear, non-protein ligands (2 CN-, 1 CO) coordinate the Fe atom in the active center (Happe, 1997; Higuchi et al., 1997; Pierik et al., 1999).

[FeS] clusters of the small subunit of [NiFe] hydrogenase from D. gigas are arranged in one line with the two low potential [4Fe-4S] clusters at the proximal and distal sides and the high potential [3Fe-4S] cluster in the middle. It was suggested that an electron channel is formed from the center of the protein, where H2 is oxidized at the active site, to the surface of the [NiFe] hydrogenase, where the electrons would be accepted by cytochrome c3. However, the role of the median [3Fe-4S] cluster is uncertain, considering the high potential of this cluster for the electron transfer is unfavorable. To study the role of the [3Fe-4S] cluster in the electron transfer, the [3Fe-4S] cluster was converted to a [4Fe-4S] cluster by site-directed mutagenesis in [NiFe] hydrogenase from D. fructosovorans. Because the catalytic activities of this mutant were similar to those of the wild-type, it was speculated that the [3Fe-4S] cluster may serve a structural function rather than participate in electron transfer (Rousset et al., 1998b). Studies by Higuchi et al. (1994) demonstrated the presence of three [FeS] clusters and one Ni ion in the [NiFe] hydrogenase of D. vulgaris strain Miyazaki F, suggesting a similar structure as the one for D. gigas [NiFe] hydrogenase. The first [NiFeSe] hydrogenase in sulfatereducing bacteria was recognized by Rieder et al. (1984) in Desulfomicrobium norvegicum, formerly Desulfovibrio desulfuricans strain Norway 4 (Sharak Genthner et al., 1997). The coding genes for the small and large subunit of the [NiFeSe] hydrogenase exhibited much sequence similarity with the corresponding genes of [NiFe] hydrogenase from Desulfovibrio gigas (Voordouw et al., 1989a). The large subunit of [NiFeSe]

CHAPTER 1.22

hydrogenase contains equimolar amounts of selenium and nickel. Another [NiFeSe] hydrogenase was purified from Desulfovibrio salexigens (Teixeira et al., 1986). Spectroscopic studies suggested that selenocysteine takes part in the coordination of the active-site nickel ion in the [NiFeSe] hydrogenase of Desulfomicrobium baculatum (Eidsness et al., 1989), formerly Desulfovibrio baculatus (Rozanova et al., 1988a). Comparative studies suggest that the [NiFeSe] hydrogenases are distinct from [NiFe] hydrogenases in terms of catalytic properties (Teixeira et al., 1987). The mechanism of selenium incorporation into proteins has been well studied with formate dehydrogenase from Escherichia coli. Selenium is present in proteins as selenocysteine, the 21st amino acid, which is cotranslationally incorporated into the nascent polypeptide from selenocysteyl-tRNASec. This selenocysteyltRNASec is synthesized from seryl-tRNASec and selenophosphate by selenocysteine synthase (Böck et al., 1991; Heider and Böck, 1993). Selenocysteyl-tRNASec recognizes an in-frame UGA codon that otherwise terminates translation (Leinfelder et al., 1988). Efficient readthrough of the UGA codon is dependent on a specific secondary structure of the mRNA downstream of the UGA codon (Zinoni et al., 1990). The selenocysteine-loaded tRNASec is directed to the UGA codon by a specialized elongation factor, SelB (Baron et al., 1993). The corresponding triplet was identified in the sequence of the coding gene for [NiFeSe] hydrogenase from Desulfomicrobium baculatum (Menon et al., 1987; 1993). The gene coding for the selenocysteineinserting tRNASec (selC) was cloned and sequenced from Desulfomicrobium baculatum (Tormay et al., 1994). A lacZ-fusion of the gene coding for the large subunit of the [NiFeSe] hydrogenase from Desulfomicrobium baculatum was constructed to study its heterologous expression in E. coli. Interestingly, in E. coli, selenocysteine was not incorporated into the D. baculatum hydrogenase subunit, demonstrating that the UGA codon was suppressed. Gel-shift experiments showed that purified SelB from E. coli in comparison to that from D. baculatum had a lower affinity for the hydrogenase mRNA from D. baculatum. Thus it appears that the specific interaction of SelB and target mRNA is a prerequisite for proper synthesis of the selenoprotein (Tormay and Böck, 1997). First evidence for a periplasmic location of a [NiFe] hydrogenases came from purification procedures that only required cells to be washed in slightly alkaline solution (Bell et al., 1974). Further studies on the localization revealed that hydrogenase in various sulfate-reducing bacteria are often localized in the periplasm. Investigated

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

species are Desulfovibrio vulgaris strain Marburg (Badziong and Thauer, 1980), Desulfovibrio vulgaris strain Hildenborough (van der Westen et al., 1978), Desulfovibrio desulfuricans (Steenkamp and Peck, 1981), Desulfovibrio gigas (Bell et al., 1974), Desulfomicrobium norvegicum (Rieder et al., 1984). Association of hydrogenases with the cytoplasmic membrane was demonstrated in Desulfovibrio vulgaris by means of immunocytochemical labeling and electron microscopy. In this species, the [NiFe] hydrogenase was located on the periplasmic side and the [NiFeSe] hydrogenase on the cytoplasmic side of the membrane (Rohde et al., 1990). In the case of Desulfovibrio desulfuricans (Essex 6) and the Gram-positive Desulfotosporosinus orientis, a cytoplasmic location of hydrogenase has been demonstrated by the use of inhibiting agents (Cypionka and Dilling, 1986a; Fitz and Cypionka, 1989). In the case of periplasmic hydrogenases, an export mechanism for these enzymes must exist. Indeed, the gene for the small subunit of the [Fe] hydrogenase of D. vulgaris was shown to encode a protein with a signal peptide of 34 amino acids (Prickril et al., 1986). There is, however, no evidence for a leader sequence in the gene for the large subunit. The situation is similar in the case of the periplasmic [NiFe] hydrogenase of Desulfovibrio gigas and of the [NiFeSe] hydrogenase of Desulfomicrobium baculatum. The mature small-subunit sequences are preceded by N-terminal signal sequences of 32 and 50 amino acids, respectively, whereas no leader sequences were found for the large subunits (Voordouw et al., 1989a; Menon et al., 1987). Also the small subunit of [NiFe] hydrogenase from Desulfovibrio desulfuricans contains a signal peptide with 50 amino acids (Rousset et al., 1990). The presence of an internal signal sequence in the large subunit of the D. vulgaris hydrogenase that might be involved in the translocation of the protein to the periplasm has been the subject of speculation. Alternatively, the immature small subunit might function as a carrier for the large subunit in the translocation process (Prickril et al., 1986). Some evidence for the latter model was presented by van Dongen et al. (1988). Homology studies revealed a consensus box containing two consecutive arginine residues in the N-terminal leader sequence of the small subunit of hydrogenases (Voordouw, 1992; Berks, 1996). A similar export mechanism was suggested for the periplasm-orientated HydB subunit of the membrane integral [NiFe] hydrogenase from the sulfur reducer Wolinella succinogenes (Gross et al., 1999). Fusion of the signal peptide from [NiFe] hydrogenase of Desulfovibrio vulgaris (Hildenborough) to the blactamase from Escherichia coli lacking its own

697

leader sequence allowed export of the enzyme. Exchange of one of the two arginines in the leader sequence to glutamate by site-directed mutagenesis inhibited export of b-lactamase completely (Nivière et al., 1992). These results demonstrated an essential role of the two subsequent arginines of the consensus box in the export of hydrogenase (Berks, 1996). The large subunit of the [NiFe] hydrogenase from Desulfovibrio gigas was shown to be processed by cleavage of 15 amino acids from the carboxy terminus (Menon et al., 1993). Hatchikian et al. (1999) demonstrated that also the large subunit of [Fe] hydrogenase from Desulfovibrio desulfuricans is subjected to a C-terminal processing in which 24 amino acids are cleaved. This finding is in agreement with the structural analysis of the same enzyme (Nicolet et al., 1999). Hatchikian et al. (1999) speculated that the C-terminal processing may play a role in the export of the protein to the periplasm. Export of the [NiFe] hydrogenase from Desulfovibrio fructosovorans may employ yet another mechanism involving an additional protein. Downstream of the structural hynA and hynB genes a third open reading frame (hydC) was identified. All three genes were found to constitute a single operon with a strong (70-like promoter. The HydC protein possesses an amphipathic segment and is speculated to mediate the integration of hydrogenase into the membrane or the export of the enzyme to the periplasm (Rousset et al., 1993). Primary acceptors for the electrons produced by hydrogenase is the periplasmic cytochrome c3 which contains multiple heme groups. Initial indication for the electron transfer between hydrogenase and cytochrome c3 arose from the co-localization of the two proteins in the periplasm (Bell et al., 1974). The interaction between hydrogenases and cytochrome c3 has been demonstrated with [Fe] hydrogenase from Desulfovibrio vulgaris strain Hildenborough (Brugna et al., 1998), [NiFe] hydrogenase from Desulfovibrio gigas (Moreno et al., 1993) and [NiFeSe] hydrogenase from Desulfovibrio desulfuricans strain Norway (Haladjian et al., 1991). The structural analysis of the cytochrome c3 molecules from Desulfovibrio vulgaris strain Hildenborough (Matias et al., 1993), Desulfovibrio desulfuricans strain Norway (Czjzek et al., 1994) and Desulfovibrio gigas (Fritz et al., 1999) revealed an overall similar molecular structure and arrangement of the four heme groups. Two types of interactions were identified, one between hemes I and II and another between hemes III and IV. A point mutation of the tyrosine 73 residue in cytochrome c3 from Desulfovibrio desulfuricans (Norway) resulted in a change of the heme IV environment and an

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R. Rabus, T.A. Hansen and F. Widdel

alteration of the hydrogenase-cytochrome interaction (Aubert et al., 1997; Aubert et al., 1998a). The positive charges surrounding the surfaceexposed heme IV of cytochrome c3 are supposed to mediate the contact to hydrogenase. In Desulfovibrio vulgaris (Hildenborough), further transfer of electrons is assumed to proceed from cytochrome c3 to the 16 heme containing highmolecular-mass cytochrome c, termed Hmc (Pollock et al., 1991; Bruschi et al., 1992; Voordouw, 1995; Pereira et al., 1998). The Hmc, which is localized to the periplasmic aspect of the membrane, is part of a multisubunit protein complex that contains membrane integral components (Rossi et al., 1993). Electrons from reduced Hmc are proposed to be transferred via the membrane integral subunits of the Hmc complex to the [Fe-S] cluster-containing gene product of Orf6 that is also part of the Hmc complex and is located at the inner aspect of the membrane. Further transfer of electrons may proceed directly to APS reductase or sulfite reductase or may involve cytoplasmic electron carriers such as flavodoxin (Voordouw, 1995). Mutants of Desulfovibrio vulgaris (Hildenborough) that had an elevated expression of the hmc operon grew more rapidly than the wildtype on H2, supporting the involvement of the Hmc complex in the electron transfer from H2 to sulfate (Keon et al., 1997). In Desulfovibrio gigas, Hmc was shown to accept electrons directly from hydrogenase (Chen et al., 1994a). Similarily, the [NiFe] hydrogenase from Desulfovibrio desulfuricans can reduce the Hmc-analogous nonaheme cytochrome c in addition to tetraheme cytochrome c3 (Fritz, 1999). These findings imply that cytochrome c3 may not always be required as a connecting link for the electron transfer from periplasmic hydrogenase to membrane-localized Hmc complex. The finding of periplasmic hydrogenase in sulfate-reducing bacteria led to the hypothesis of energy conservation by so-called vectorial electron transport, the simplest transmembrane process that can generate a proton gradient for chemiosmotic ATP synthesis. The protons from H2 oxidation are released by hydrogenase into the periplasm, while abstracted electrons are transported via redox-active centers of transmembrane proteins to the cytoplasm (or cytoplasmic aspect of the membrane) and used for sulfate reduction. This charge separation, which is driven by the exergonic process of sulfate reduction, is compensated by a simultaneous (somewhat “retarded”) proton flow via ATPase into the cytoplasm; ATPase finally conserves the energy from the redox process in a phosphoric anhydride bond. The generation of a proton gradient by simple charge separation (vectorial electron transport) by a periplasmic hydroge-

CHAPTER 1.22 + + +

– – –

2 H+ SO42– H+

21/3 ATP

7 H+

10 H+

H2 ase 4 H2

SO42–

8 e–

2 ATP

S R

x H+

2 ADP + 2 Pi

x / ATP 3

H2S

x H+ H2S + + +

– – –

Fig. 10. Possible generation of a proton-motive force (pmf) during growth of Desulfovibrio on H2 and sulfate (low concentration). Electrogenic transport of sulfate with three protons is assumed (Cypionka, 1989). In addition to a proton-translocating mechanism during sulfate reduction, vectorial electron transport from a periplasmic hydrogenase (H2ase) via the membrane may contribute to the generation of a pmf. Periplasmic cytochrome c3 and membrane– spanning, high-molecular-mass cytochrome (Hmc) mediate the electron flow between H2 oxidation and sulfate reduction. Activation of sulfate consumes 2 ATP because AMP liberated by the adenosine-5¢-phosphosulfate (APS) reductase from APS has to be converted to ADP by adenylate kinase (myokinase). Abbreviation: SR, enzymes and other components involved in sulfate reduction to H2S.

nase would leave eight extracellular protons per mol sulfate to be reduced. Because (at least) one of these electrogenically produced protons enters the cell during sulfate-transport (Cypionka, 1989), no more than seven protons would be left for chemiosmotic energy conservation yielding 13/4 to 21/3 mol ATP, if one assumes a ratio of 3–4 H+/ATP (Schink, 1988a; Thauer and Morris, 1984; Stock et al., 1999) per mol sulfate reduced with H2 (Fig. 10). Because sulfate activation is associated with a net consumption of 2

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

ATP/SO42-, a maximum of 1/3 mol ATP would remain for cell synthesis. This is much less than the estimates from growth yields, which suggest a net synthesis of 1.3 mol ATP per mol sulfate (Nethe-Jaenchen and Thauer, 1984). Hence, a proton gradient seems to be generated in addition by proton pumping, provided the 3H+/ATP ratio used in the calculations is a correct estimate. Indeed, proton translocation with H2 and sulfate has been measured in Desulfovibrio desulfuricans strain Essex 6 in which the hydrogenase present under the applied growth conditions was reported to be cytoplasmic or at least on the cytoplasmic aspect of the membrane (Fitz and Cypionka, 1989). Strains of other Desulfovibrio species translocated protons with H2 and nitrite, even though hydrogenase and nitrite reductase were both periplasmic enzymes (Barton et al., 1983; Steenkamp and Peck, 1981); this location excludes generation of a proton gradient by simple vectorial electron flow via the membrane. Finally, growth of the Gram-positive Desulfotosporosinus orientis on H2 with high cell yields demonstrated that chemiosmotic ATP synthesis does not require a periplasmic hydrogenase (Cypionka and Pfennig, 1986). Hence, vectorial electron transport due to periplasmic hydrogenase appears to be only an additional mechanism for energy conservation in a number of Desulfovibrio species. The main mechanism is obviously vectorial proton transport (e.g., by proton-pumping redox proteins or “Mitchell-type” loops, involving the menaquinones that are commonly present in sulfate-reducing bacteria). Nothing is known about the possibility of a Q-cycle (Peck and Lissolo, 1988); considering this translocates two protons for one electron, the process would require significant differences in the redox potential between two couples, which is not very likely in the anaerobic respiratory chain in sulfate-reducing bacteria. An energetically intriguing, not sufficiently understood, aspect is the growth of sulfatereducing bacteria with H2 over a wide range of partial pressures. At standard pressure, H2 is one of the energetically most favorable electron donors (DG∞¢ = –152.2 kJ/mol sulfate). However, Desulfovibrio was shown to scavenge H2 below 10 Pa (10-4 atm; Cord-Ruwisch et al., 1988). In natural anoxic habitats where sulfate-reducing bacteria thrive, even H2 partial pressures as low as >5 Pa (>5 · 10 -5 atm; Sørensen et al., 1981) 2.5 · 10 -2 Pa (2.5 · 10 -7 atm, would be at the thermodynamic equilibrium; Scranton et al., 1984) and 1.1 Pa (1.1 · 10 -5 atm; Lovley et al., 1982) have been measured. The free energy of sulfate reduction with H2 at varying partial pressure is depicted in Fig. 11. Assuming that net ATP synthesis coupled to any catabolic overall

699

50 2– 4

CHAPTER 1.22

0



2–

–50

H2

f

SO 4

f

2 4

–100   G

–150 –200 10 –3 10 –2 10 –1

1

101 10 2 10 3 10 4 10 5 (Pa)

10 8 10 7 10 6 10 5 10 4 10 3 10 2 10 1 (

1



H2 Fig. 11. Free energy change of sulfate reduction and “reverse” methanogenesis related to H2 partial pressure. Reduction of SO42- with H2 is shown in red line; H2 formation from CH4 is shown in blue lines. Full lines represent values calculated for pH 7 and dashed lines those for pH 8.

reaction has irreversible character and requires around 70 kJ/mol ATP, the ATP gain at the natural, low H2 pressures has to be much less than the gain measured with optimal supply of the electron donor. Hence, electrons from H2 at low partial pressure cannot be transported via a chain with the same number of energy-conserving steps (“coupling sites”) as electrons from H2 at standard pressure. One may speculate that electrons from H2 at various partial pressures enter the electron transport chain at different levels, and that different types of hydrogenases are involved. Formate Formate is a fermentation product in several anaerobic bacteria, as for instance in enterobacteria. In addition, formate has been discussed as a means for an interspecies transfer of reducing equivalents and as an alternative to H2 in natural anaerobic bacterial communities (Thiele et al., 1988a; Thiele and Zeikus, 1988b); formate transfer was most likely to occur in a sulfate-reducing coculture (Zindel et al., 1988). However, syntrophisms based on interspecies H2 transfer are more important (Schink, 1997). Also energetically, formate may be regarded as an electron donor that is equivalent to H2. The redox potential of the couples 2 H+/H2 and HCO3-/HCOO- are very similar (E0¢ around –0.41 V). Hence, formate is a favorable electron donor: -

2-

-

4HCO2 + SO4 + H + Æ 4HCO3 + HS DGo ¢ = -146.6 kJ mol sulfate

(20)

700

R. Rabus, T.A. Hansen and F. Widdel

The ability to grow with formate has been observed in most genera of sulfate-reducing eubacteria. Formate dehydrogenase has been found in Desulfovibrio (Fauque et al., 1991; LeGall and Fauque, 1988) and in completely oxidizing sulfate reducers except for Desulfobacter (Schauder et al., 1986; Spormann and Thauer, 1988; Aeckersberg et al., 1991; Rabus et al., 1993; Fukui et al., 1999). Formate dehydrogenase in Desulfovibrio is a periplasmic protein (Odom and Peck, 1981a). It was partially purified from Desulfovibrio vulgaris (Miyazaki); purified cytochrome c553 functioned as an electron acceptor but cytochrome c3 did not (Yagi, 1979). The formate dehydrogenase of Desulfovibrio gigas is thought to use cytochrome c3 as electron acceptor (RiedererHenderson and Peck, 1986). The periplasmic formate dehydrogenase of Desulfovibrio vulgaris (Hildenborough) was purified by Sebban et al. (1995). The enzyme is composed of three subunits. The large 83.5-kDa subunit contains a molybdenum cofactor and most likely presents the active site. A 27-kDa subunit with an [Fe-S] center is similar to the [Fe-S]-containing subunit of the formate dehydrogenase from Escherichia coli. The 14-kDa subunit holds a c-type heme. Cytochrome c553 is thought to be the natural electron acceptor of this formate dehydrogenase (Sebban-Kreuzer et al., 1998b). Recently, a tungsten-containing formate dehydrogenase was purified from Desulfovibrio gigas and characterized. This protein was found to have a heterodimeric structure (subunits 92 kDa and 27 kDa) and to contain approximately seven Fe per molecule most probably in two [4Fe-4S] clusters; the tungsten is most likely bound to a molybdopterin guanine dinucleotide-type cofactor (Almendra et al., 1999). This is the second W-protein that has been isolated from Desulfovibrio gigas (see dissimilation of ethanol). The formate dehydrogenase of Desulfovibrio desulfuricans was also found to contain molybdenum, iron-sulfur centers and heme (Costa et al., 1997). Completely oxidizing sulfate-reducing bacteria employ the C1/CO dehydrogenase pathway for acetyl-CoA oxidation, always contain formate dehydrogenase (see next section) and often grow on formate. Growth of the completely oxidizing Desulfotomaculum acetoxidans on formate is poor and difficult to achieve (Widdel and Pfennig, 1981b), despite the high formate dehydrogenase activity (Spormann and Thauer, 1988). This can be explained by the lack of a proper transport system. Formic acid is less lipophilic and has a lower pKa value (3.75) than acetic acid (4.75) and probably cannot enter the cell by diffusion via the mem-

CHAPTER 1.22

brane. Formate dehydrogenases that are part of the C1/CO-pathway were found to be membrane-associated, probably with the reactive site toward the cytoplasm. Their natural electron acceptor is not known. The reduction of NAD+ with formate probably occurred via a transhydrogenase. Terminal Oxidation and Utilization of Acetate The oxidation of organic substrates in sulfatereducing bacteria may be complete, leading entirely to CO2, or incomplete with acetate being the end product; in the latter case a mechanism for acetyl-CoA oxidation is lacking. A complete oxidation of organic compounds by sulfate reducers without the capacity for acetate oxidation is possible only with C1-compounds such as formate or methanol (Klemps et al., 1985; Nanninga and Gottschal, 1987; Ollivier et al., 1988), or with C2-compounds that are more oxidized than acetate (e.g., glycine; Stams et al., 1985), glycolate (Friedrich and Schink, 1995) or oxalate (Postgate, 1963). The capacity for complete oxidation of various organic substrates, viz. the presence of a pathway for acetyl-CoA oxidation, usually includes also the ability to use free acetate as a growth substrate: 2-

-

CH3 COO- + SO4 Æ 2HCO3 + HS (21) DGo ¢ = -47.6 kJ mol sulfate Desulfobacter species use acetate preferentially or even exclusively. In certain complete oxidizers, however, growth on acetate may be very poor, even though other compounds are readily oxidized to CO2. Complete oxidizers may excrete acetate if growing, e.g., on ethanol or butyrate (Imhoff-Stuckle and Pfennig, 1983; Laanbroek et al., 1984; Schauder et al., 1986; Widdel and Pfennig, 1981b). With limiting amounts of substrates, the excreted acetate may be oxidized further. Species using acetate very poorly may leave the acetate once formed almost untouched (Imhoff-Stuckle and Pfennig, 1983). An explanation for the acetate excretion by complete oxidizers is that formation of acetyl-CoA proceeds faster than its terminal oxidation. The formation of 1 mol acetate per mol butyrate oxidized has been explained by the use of 1 mol acetyl-CoA (from 2 mol formed per mol of butyrate) for the activation of butyrate by a CoA transferase (Schauder et al., 1986); see also section “Butyrate and other fatty acids”). The marginal capacity or inability of some complete oxidizers to use free acetate is not clearly understood.

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

701

Acelate ATP ADP + P1 ADP ATP + P1 X[H]2

AcCoA MADH

Citr

OxAc

MAD(P)H

Citr

OxAc

MAD+

X Icitr

Mal

Icitr

Mal

MADPH

AcCoA

CO2 Z-OCl Fd(ox) SucCoA Fd(red)

Fum MKH2

MK

Succ

Acelate

Mal

Icitr

MADP+

MADP+ H2O

Citr

OxAc

MAD(P)+

MADPH

AcCoA

H2O

CO2 Fum

MKH2

MK

2-OGl Fd(ox) SucCoA Fd(red)

Succ

CO2

A

Acelate

B

MADP+ MADPH H2O

CO2 2-OGl Fd(ox) SucCoA Fd(red)

Fum X[H]2 X

Succ

CO2

CO2 ATP ADP +P1 C

Fig. 12. Modifications of the citric acid cycle for the anaerobic oxidation of acetate in three species of sulfate- and sulfurreducing bacteria: (A) Desulfobacter postgatei; (B) Desulfuromonas acetoxidans; and (C) Desulforella acetivorans. The reactions leading from citrate to succinyl-CoA are the same in all three cycles. The H+ ions, water, and some other reactants are not indicated. Abbreviations: AcCoA, acetyl-CoA; Citr, citrate; Fd(ox), oxidized ferredoxin; Fd(red), reduced ferredoxin; Fum, fumarate; Icitr, isocitrate; Mal, malate; MK, menaquinone; 2-OGl, 2-oxoglutarate (a-ketoglutarate); OxAc, oxaloacetate; Succ, succinate; SucCoA, succinyl-CoA; X, unknown physiological electron or hydrogen carrier. Adapted from Thauer (1988) and Thauer et al. (1989b).

Organic end products other than acetate are formed in incomplete or complete oxidizers if substrate oxidation leads to products that cannot be degraded further by the enzymatic outfit. Examples are the oxidation of n-propanol, nbutanol or isobutanol to propionate, butyrate or isobutyrate, respectively (Mechalas and Rittenberg, 1960), the formation of propionate from C-odd fatty acids (Pfennig and Widdel, 1981b; Widdel, 1980; Widdel and Pfennig, 1981b) or of benzoate from phenylpropionate (Brysch et al., 1987). The pathways for acetyl-CoA oxidation have been elucidated by enzymatic measurements and growth experiments with 14C-labeled substrates. In Desulfobacter postgatei, all enzymes of a citric acid cycle were found (Brandis-Heep et al., 1983; for an overview see Kröger et al., 1988; Thauer, 1988; Thauer, 1989a; Thauer et al., 1989b). Also with position-labeled [14C]-acetate as growth substrate, the labeling pattern in aspartate and glutamate that are derived from oxaloacetate and 2-oxoglutarate were in agreement with an operating citric acid cycle (Gebhardt et al., 1983). The cycle differs in some aspects from the cycles in mitochondria and aerobic bacteria. Acetate in Desulfobacter is not activated via acetate thiokinase (acetyl-CoA synthetase) or acetate kinase and phosphotransacetylase (phosphate acetyltransferase), but via

succinyl-CoA:acetate CoA transferase (Fig. 12). Dehydrogenation of isocitrate occurs with NADP+, as in most eubacteria. However, the conversion of 2-oxoglutarate to succinyl-CoA does not couple to NAD+, but rather to a ferredoxin, as electron acceptor. The hydrogen acceptor for succinate oxidation to fumarate, so far known, is menaquinone and not ubiquinone, as in mitochondria and most Gram-negative bacteria. A remarkable finding was that condensation of acetyl-CoA and oxaloacetate to citrate in Desulfobacter is associated with ATP synthesis (Möller et al., 1987). The enzyme, ATP-citrate lyase, enables the conservation of the energy of the thioester; the citrate synthase reaction in the common citric acid cycle wastes this energy by hydrolysis of the intermediary citryl-CoA. The ATP-citrate lyase reaction is reversible (DG∞¢ (0 kJ). Indeed, before being found in Desulfobacter, the reaction was only known to proceed in vivo in its opposite direction. In the cytosol of eukaryotic cells, ATP-citrate lyase cleaves citrate that functions as the acetyl carrier across the two mitochondrial membranes. Green sulfur bacteria fix CO2 via a reverse citric acid cycle which was found to include the ATP-citrate lyase reaction (Ivanovsky et al., 1980). Citrate formation in Desulfobacter species occurs with Si-face stereospecificity. The acceptor for malatedehydrogenase is neither NAD+ nor NADP+. The reduction of the artificial acceptor 2,6-

702

R. Rabus, T.A. Hansen and F. Widdel

dichlorophenol indophenol (DCPIP) was inhibited by the menaquinone antagonist 2(nheptyl)-4-hydroxyquinoline-N-oxide (HQNO). From this and the occurrence of the activity in the membrane, one may speculate that menaquinone serves as hydrogen acceptor in malate oxidation. However, in vitro tests with substitutes for menaquinone (naphthoquinone, dimethylnaphthoquinone) yielded no or marginal activity (Möller-Zinkhan and Thauer, 1988). Still, it is most likely that Desulfobacter employs a more positive acceptor for malate oxidation than other bacteria. The reversible, energy-conserving ATP-citrate lyase reaction in Desulfobacter necessitates the use of a more positive acceptor for malate oxidation to favor the concentration of the product. The citric acid cycle in Desulfobacter hydrogenophilus has the same reactions as in D. postgatei (Schauder et al., 1987). A comparison of the modifications of the citric acid cycle found in sulfate- and sulfurreducing bacteria is presented in Fig. 12. In completely oxidizing sulfate reducers other than Desulfobacter, 2-oxoglutarate dehydrogenase could not be detected (Schauder et al., 1986). Inasmuch as most completely oxidizing sulfate reducers grow very poorly on acetate, [3-14C]-pyruvate was used for them as growth substrate in labeling studies (Schauder et al., 1986). For Desulfotomaculum acetoxidans, [14C]acetate could be used. The labeling in aspartate and glutamate showed that a citric acid cycle was not operating. Citrate synthase of the incomplete cycle seemed to have re-specificity. All complete oxidizers without 2-oxoglutarate dehydrogenase contained high activities of CO dehydrogenase, which was absent in Desulfobacter species. In labeling experiments, cell extracts of species without 2-oxoglutarate dehydrogenase catalyzed an equilibrium exchange of the C1-position in acetyl-CoA with free CO2. Furthermore, these species formed traces of methane indicating a reactive methyl group as an intermediate; such a mini-methane formation was not observed in Desulfobacter. All these findings led to the conclusion that completely oxidizing genera other than Desulfobacter, viz. the majority of sulfate reducers, cleave acetyl-CoA into bound CO and a bound methyl group; both C1 units are then oxidized to CO2 (Schauder et al., 1986; Spormann and Thauer, 1988; Fig. 13). The carrier of the methyl group in Desulfotomaculum acetoxidans was tetrahydrofolate (Spormann and Thauer, 1988). For two steps, namely the dehydrogenation of methylenetetrahydrofolate and formate, NAD+ wasthe natural electron acceptor. The conversion of formyl-tetrahydrofolate to free formate is associated with ATP synthesis. In Desulfobacterium autotrophicum, the C1-carrier is a homologue of tetrahydrofolate, tetrahy-

CHAPTER 1.22

dropteroyltetraglutamate, which has four glutamate residues instead of one (Länge et al., 1989). Dehydrogenation of the methylene group in this species occurs with NADP+ (Schauder et al., 1989). The results demonstrated for the first time that the pathway of acetyl-CoA synthesis known from homoacetogenic bacteria (Fuchs, 1986; Wood et al., 1986) can operate in the reverse direction for terminal oxidation of organic substrates. Thereafter, the pathway was also found in a syntrophic thermophile that oxidized acetate to CO2 and H2 of low partial pressure (Lee and Zinder, 1988), and in the archaeal sulfate reducer, Archaeoglobus (Möller-Zinkhan et al., 1989). The bioenergetic implications of the pathways for acetate oxidation and terminal oxidation of other organic compounds in sulfate- and sulfurreducing bacteria have been discussed (Kröger et al., 1988; Thauer, 1988; Thauer et al., 1989b). The free energy gain from reduction with acetate (equation 21) is much lower than from sulfate reduction with H2 at standard pressure (Fig. 11). However, due to the stoichiometric factors in the equations, the free energy per mol of sulfate reduced is less concentration-dependent in the case of acetate oxidation than in the case of hydrogen oxidation. Net ATP yields available for cell synthesis may be estimated from growth yields. The highest growth yield measured with an acetate-oxidizing sulfate reducer, Desulfobacter postgatei, in batch cultures was 4.8 g dry mass/mol acetate (or sulfate). Theoretical maximum growth yields (Ymax) from extrapolation to infinite growth rates (m = µ) have not been determined. Nevertheless, the growth yield may be compared to that of other bacteria at similar growth rates. The doubling time of D. postgatei was around 20 hours. At this doubling time, Desulfovibrio vulgaris growing on H2 and sulfate with acetate as carbon source for cell synthesis would have a growth yield of 7.7 g dry weight/mol sulfate (calculated from Nethe-Jaenchen and Thauer, 1984). A yield of 1.3 mol ATP/mol sulfate was estimated for D. vulgaris (Nethe-Jaenchen and Thauer, 1984). Desulfobacter postgatei should thus gain around 0.8 mol ATP/mol acetate. Comparison of the free energy available from reactions (21) with the generally observed requirement of >70 kJ/mol ATP (Schink, 1988a; Thauer et al., 1977) again indicates that acetateoxidizing sulfate-reducing bacteria obtain less than 1 mol ATP/mol sulfate. In Desulfobacter, ATP-citrate lyase enables net gain of 1 ATP by substrate-level phosphorylation during acetate oxidation; however, 2 ATP are needed for sulfate activation (assuming that pyrophosphate is irreversibly hydrolyzed). The energy requirement for sulfate transport in

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes Organic electron donor

703

Organic electron donor

O

— —

— —

O

— C— — SCoA CH3 —

— SCoA — C— CH3 —

CH3 — THP

[CO]

[CO]

CH3 — THMP

X X[H]2

F420

Quinonered

F420H2

Quinoneox

— THMP CH2 —

— THP CH2 — NAD(P)+ NAD(P)H X

F420

Quinonered

F420H2

Quinoneox

— THMP CH —

— THP CH —

X

X[H]2

X[H]2

CHO — THMP

CHO — THP ADP + Pi ATP

CHO — MF

–

HCOO

X

NAD+

X[H]2

NADH CO2

CO2

A

CO2

CO2

B

Fig. 13. Terminal oxidation of acetyl-CoA via the C1/carbon monoxide dehydrogenase pathway in sulfate-reducing bacteria. The H+ ions, water, and some other reactants are not indicated. (A) Reactions in Desulfotomaculum acetoxidans, Desulfobacterium autotrophicum, and presumably other completely oxidizing sulfate-reducing bacteria (except for Desulfobacter species). The former species uses NAD+, and the latter NADP+ for dehydrogenation of the methylene group; in Desulfotomaculum, NAD+ is probably not the direct electron acceptor for formate dehydrogenase but reduced via an unknown, primary acceptor. THP is tetrahydrofolate in D. acetoxidans, and tetrahydropteroyltetraglutamate in D. autotrophicum. (B) Reactions in the archaeon Archaeoglobus fulgidus. Abbreviations: [H], unknown physiological electron or hydrogen carrier; MF, methanofuran; THMP, tetrahydromethanopterin; THP, tetrahydropterin; F420, formate dehydrogenase. Adapted from Thauer (1988) and Thauer et al. (1989b).

Desulfobacter is unknown. Desulfobacter occurs mainly in brackish or marine environments with high sulfate concentrations. It is therefore likely that an electrogenic transport of sulfate (Warthmann and Cypionka, 1990) is not required under such conditions. Thus, for a net ATP gain, more than 1 ATP has to be synthesized by chemiosmosis. In Desulfuromonas, there is no substrate-level phosphorylation. In Desulfobacter postgatei, the transport of reducing equivalents from NADPH to MK-7 (DE0¢ = 0.25 V) could be associated with a translocation of 2 H+/2 [H], which are 4 H+/acetate (Kröger et al., 1988; Thauer, 1988). The preceding reduction of NADP+ with ferredoxin (DE0¢ = 0.1 V) has been discussed as another energy-conserving step allowing the translocation of 1 H+ (Fig.

14A). With the assumed requirement of 3–4 H+/ ATP, chemiosmosis in Desulfobacter postgatei should produce 5/4 to 5/3 ATP and thus allow a net ATP gain of 1/4 to 2/3 per sulfate for cell synthesis. The latter value is more likely in view of the aforementioned calculations based on growth yields. In Desulfobacter, succinate oxidation to fumarate (E0¢ = +0.033 V) with menaquinone (E0¢ = –0.074 V) is endergonic from the viewpoint of standard potentials. Still, the reaction appears possible with shifted concentration ratios of involved redox couples, or by specific coupling to a favorable redox reaction of the sulfate reduction pathway. It is true that from a mere thermodynamic viewpoint any unfavorable partial reaction is rendered possible if embedded in an exergonic overall reaction. In biological systems,

704

R. Rabus, T.A. Hansen and F. Widdel

E¢ (V)

–0.4

CHAPTER 1.22

2-OGl

2-OGl

Fd(red)

Fd(red)

Icitr

~ +? ∆µ H

[CO]

HCOO– ~ +? ∆µ H

Icitr NADH

NADPH

–0.3

~ +? ∆µ H

NAD NADPH

CH2 =THF ~ +? ∆µ H

~ +? ∆µ H

–0.2 Mal ?

Y Mal

[S]

X

HSO3–

–0.1 MKH2

MKH2

HSO3–

CH3 =THF MKH2

APS

0.0 Succ

APS

Succ

A

B

C

Fig. 14. Flow of reducing equivalents during terminal oxidation of acetyl-CoA in sulfate- and sulfur-reducing bacteria. Electron-donating and electron-accepting redox couples are presented only as the reduced or oxidized forms, respectively. In most cases, the midpoint potential is indicated (often concentration-independent, with E’ being identical to E 0¢); the exact redox potential in the cell may differ, according to concentrations of reaction partners. The redox potential of APS reduction refers to concentrations of 1 mM. The redox potential of bisulfite reduction given for a six-electron step refers to 1 mM HSO 3and a range of 1 to 10 mM H2S (dotted redox span). The concentration range of H2S for sulfur reduction is also 1 to 10 mM. Bound CO probably has a less negative midpoint potential than free CO. The value of the former is not exactly known. (A) Desulfobacter postgatei, growing on acetate and sulfate. (B) Desulfuromonas acetoxidans, growing on acetate and sulfur. X is a carrier, presumably a cytochrome c that donates electrons to sulfur reduction. (C) Desulfotomaculum acetoxidans, growing on acetate and sulfate. Y is an unknown electron carrier. Symbols and abbreviations: arrows (in full lines), reactions catalyzed by membrane-associated enzymes; arrows (in dashed lines), reactions catalyzed by soluble enzymes; APS, adenosine-5¢phosphosulfate. Fd(red), reduced ferredoxin; Icitr, isocitrate; Mal, malate; Succ, succinate; THF, tetrahydrofolate; 2-OGl, 2oxoglutarate. Adapted from Möller-Zinkhan and Thauer (1988) and Thauer (1988).

however, also the rates are important. In a thermodynamically very unfavorable partial reaction, the very low product concentration may not be sufficient to allow appropriate rates with the enzyme of the subsequent reaction. The equilibrium of unfavorable reactions can in principle be shifted by an input of energy, which in case of redox reactions is known as reversed electron transport. Indeed, a membrane preparation catalyzed a strictly ATP-dependent oxidation of succinate with sulfur or NAD+ (Paulsen et al., 1986). The reaction was sensitive to the ATPase inhibitor DCCD6 or to the protonophore TTFP7, indicating that ATP acted indirectly via formation of a proton gradient as the driving force for succinate oxidation. The pri-

mary hydrogen acceptor of succinate oxidation was apparently menaquinone (MK-8); its analogue dimethylnaphthoquinone was reduced with succinate without addition of ATP, as in Desulfobacter. Hence, the proton gradientdriven reaction is probably the oxidation of menaquinone with the subsequent electron carrier that feeds into sulfur reductase. This electron carrier might be one of the membranebound c-type cytochromes with a midpoint potential more negative than –200 mV. It has been estimated that two to three protons have to reenter the cell to promote the oxidation of one molecule of succinate (Kröger et al., 1988; Thauer, 1988). With the remaining one to two protons, the net energy conservation in the sulfur

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes Lactate

Ethanol

705

Propionate ATP ADP Propionyl- P 2X

Succinyl-CoA

2X[H]2

2X

ADP + Pi ATP

Propionyl-CoA 2X[H]2 Methylmalonyl-CoA TC COO–

Succinate 2X

TC

2X[H]2 Fumarate

Oxaloacetate Acetaldehyde

Pyruvate 2X 2X[H]2, CO2 2X

2X[H]2

NADH NAD+

Malate

Acetyl-CoA

Acetyl- P ADP ATP Acetate Fig. 15. Oxidation of ethanol, lactate, and propionate to acetate in Desulfobulbus propionicus. Methylmalonyl-CoA is formed by carboxylation of propionyl-CoA with CO2 bound to transcarboxylase (TC). The “X” is the unknown physiological electron or hydrogen carrier.

reducer would be 1/3 to 2/3 ATP per molecule of acetate. Suggested proton-translocating reactions in the C1/CO-dehydrogenase-pathway are indicated in Fig. 14C. Propionate Propionate serves as electron donor and carbon source for the incompletely oxidizing Desulfobulbus species and several completely oxidizing sulfate reducers (Table 2). Propionate in Desulfobulbus is oxidized to acetate via a randomizing pathway with succinate, a symmetric molecule, as free intermediate (Kremer and Hansen, 1988a; Fig. 15). The principle of this pathway in Desulfobulbus was first elucidated in its reverse direction, the formation of propionate from fermentable substrates in the absence of sulfate (see Fermentative and Syntrophic Growth in the Absence of Sulfate). A succinate dehydrogenase/fumarate reductase was purified from Desulfobulbus elongatus; it consists of three subunits and contains one cytochrome b, flavin

and eight non-heme iron atoms (Samain et al., 1987). The oxidation of propionate to CO2 by Desulfococcus multivorans was also shown to proceed via the succinate pathway (Stieb and Schink, 1989). Unlike Desulfobulbus, Desulfococcus can oxidize the pyruvate formed via the C4-dicarboxylic acid sequence further than the acetate level; acetyl-CoA is oxidized to CO2 via the C1/CO-dehydrogenase-pathway (see preceding section). Not unexpectedly therefore, the molar growth yield of Desulfococcus, if related to propionate, was more than twice as high as that of Desulfobulbus (approximately 10 and 4 g dry mass per mol propionate, respectively; Stieb and Schink, 1989; Stams et al., 1984); if related to sulfate, the yields are rather similar (approximately 5.7 and 5.3 g dry mass per mol sulfate, respectively). Butyrate and Other Fatty Acids Butyrat and higher fatty acids are oxidized by many incompletely and completely oxidizing sulfate-

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CHAPTER 1.22

reducing bacteria (Widdel, 1980; Pfennig and Widdel, 1981b; Widdel, 1988; Table 2). The incomplete oxidation of C-even fatty acids yields only acetate. The C-odd fatty acids are oxidized to acetate and propionate. Measured degradation balances were in agreement with the following general equations: 2CH3 (CH 2 )2 n COO- + n SO4 Æ 2 (n + 1) CH3 COO- + n H 2 S 2 2CH3 (CH 2 )2 n +1 COO- + n SO4 Æ 2 nCH3 COO- + CH3 CH 2 COO- + n H 2 S 2

(13)

(14)

The ratio is explained by a b-oxidation. In the case of C-odd fatty acids, propionyl-CoA is left, which obviously cannot be metabolized by most incomplete fatty acid oxidizers and therefore has to be excreted. If 2-methylbutyrate is used by incomplete oxidizers, propionate is formed too. b-Oxidation of acetyl-CoA is in principle not hampered by a 2-methyl group; this leads to formation of propionyl-CoA rather than acetylCoA from the first part of the fatty acid chain. Most complete oxidizers can degrade the propionyl residue; therefore, also C-odd fatty acids and, if metabolized, 2-methylbutyrate are oxidized like C-even fatty acids: 2-

4H(CH 2 )n COO- + (3n + 1) SO4 + (2n + 2) H + Æ (4n + 4) HCO3 + (3n + 1) H 2 S (15) Nevertheless, complete oxidizers may excrete acetate, probably as a result of an “imbalance” between b-oxidation and acetyl-CoA oxidation. Desulfobacterium autotrophicum formed one mol acetate per mol butyrate (Schauder et al., 1986). It is concluded from this ratio that acetate was formed by CoA transfer from acetyl-CoA to activate butyrate. Free acetate is used very poorly by this sulfate reducer. Desulfotomaculum acetoxidans, which is a complete oxidizer but cannot utilize propionate, oxidizes valerate to propionate (Widdel and Pfennig, 1982). Sulfate reducers using isobutyrate and 3-methylbutyrate (isovalerate) are always complete oxidizers. The pathway of isobutyrate degradation has been elucidated in a Desulfococcus multivorans strain (Stieb and Schink, 1989). The reactions are in principle the same as found in aerobic organisms’ metabolism of valine. The initial degradation steps in Desulfococcus were mediated by two enzymes that are involved in the catabolism of n-butyrate. Isobutyryl-CoA is first converted via butyryl-

CoA dehydrogenase and enoyl-CoA hydratase to 3-hydroxyisobutyryl-CoA, which is then hydrolyzed to the free acid and oxidized to methylmalonate semialdehyde. CoA-dependent dehydrogenation of the semialdehyde and decarboxylation leads to propionyl-CoA. This is oxidized to acetyl-CoA as in Desulfobulbus propionicus (see foregoing section). Acetyl-CoA is then oxidized via the C1-pathway (Schauder et al., 1986). By means of a succinyl-CoA:acid CoA transferase, the conversion of succinylCoA to succinate is coupled to the activation of the free isobutyrate to isobutyryl-CoA. In contrast to the sulfate-reducing culture, a methanogenic coculture isomerized isobutyrate to butyrate that was oxidized to two acetate residues (Stieb and Schink, 1989). The pathway for isovalerate degradation has not been examined in sulfate-reducing bacteria. Lactate and Pyruvate Lactate, the “classical” substrate for cultivation of sulfate-reducing bacteria, is utilized by most species of almost each genus and may be oxidized completely or incompletely (equations 1 and 2, respectively). Desulfoarculus baarsii (formerly Desulfovibrio; Widdel, 1980), several Desulfobacter species (Widdel and Pfennig, 1981a; Widdel, 1987) and some species of the genera Desulfobacterium, Desulfonema (Widdel et al., 1983) and Desulfotomaculum (Widdel and Pfennig, 1981b) cannot use lactate. The oxidation of L- and D-lactate to pyruvate is mediated by NAD(P)+-independent lactate dehydrogenases that occur mainly membranebound. None of the enzymes from sulfatereducing bacteria have been purified to homogeneity. D-Lactate dehydrogenase of Desulfovibrio desulfuricans was present in the particulate fraction, and detergents were required for its solubilization (Czechowski and Rossmoore, 1980), but in Desulfovibrio vulgaris strain Miyazaki, part of the enzyme activity was soluble (Ogata et al., 1981). L-lactate dehydrogenase activities were demonstrated in a Desulfomicrobium baculatum-like strain (formerly a Desulfovibrio desulfuricans strain; Stams and Hansen, 1982), Desulfovibrio desulfuricans, Desulfovibrio gigas (Peck and LeGall, 1982b) and Desulfovibrio vulgaris (Pankhania et al., 1988). In Desulfovibrio vulgaris strains, pyruvate has been shown to be oxidatively decarboxylated to acetyl-CoA with ferredoxin or flavodoxin as electron acceptor (Suh and Akagi, 1966; Ogata and Yagi, 1986). The low potential ferredoxin I was found to be particularly active in the pyruvate:acceptor oxidoreductase reaction of Desulfovibrio gigas (LeGall and Fauque, 1988; Fauque et al., 1991). Pyruvate:ferredoxin oxi-

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

doreductase (POR) has been purified from Desulfovibrio africanus. The enzyme is a homodimer of 256 kDa and contains thiamine pyrophosphate (TPP) and three iron-sulfur clusters. Spectroscopic analysis of the activated enzyme indicated the presence of a free radical (Pieulle et al., 1995). A catalytic mechanism involving a free radical had been demonstrated before for the POR from the extremely halophilic bacterium Halobacterium halobium (Cammack et al., 1980). The gene for POR of D. africanus was cloned and the enzyme overexpressed in E. coli (Pieulle et al., 1997) so that enzyme quantities sufficient for crystallization could be obtained. The enzyme from D. africanus is the first POR to have its crystal structure determined (Chabriere et al., 1999a; Chabriere et al., 1999b; Pieulle et al., 1999a; for review see Charon et al., 1999). The substrate pyruvate is bound at the active site in the proximity of TPP. The three [4Fe-4S] clusters are located between TPP and the protein surface, indicating that this arrangement serves as the path for electron transfer within the protein. Further transfer of electrons from POR to the external ferredoxin probably requires electrostatic interactions (Pieulle et al., 1999b). In the incomplete oxidation of organic substrates, acetyl-CoA produced from pyruvate is converted to acetate by means of phosphotransacetylase and acetate kinase (Brown and Akagi, 1966; Ogata and Yagi, 1986), which allows phosphorylation of ADP to ATP. The incomplete oxidation of lactate to acetate in Desulfovibrio species gave the first hint that oxidation of organic substrates in sulfatereducing bacteria is associated with chemiosmotic energy conservation (formerly, electron transport phosphorylation) in addition to substrate-level phosphorylation. A simple yet basic calculation (Peck, 1966) revealed, that the net ATP gain by substrate-level phosphorylation during growth of Desulfovibrio on lactate and sulfate is zero. The two molecules of lactate oxidized per molecule of reduced sulfate (equation 1) yield two molecules of ATP during liberation of acetate via acetate kinase; these two ATP molecules are consumed for the activation of sulfate, namely one for the ATP sulfurylase reaction and one for regeneration of ADP from AMP (adenylate kinase reaction) formed during APS reduction (see Activation of Sulfate in this Chapter). Hence, there has to be an additional mechanism for ATP formation. A unique mechanism for generation of a proton gradient with lactate as electron donor was suggested by Odom and Peck (1981b). Their socalled hydrogen-cycling model for growth on lactate of a Desulfovibrio involved the cytoplasmic production of H2 as a result of the oxidation of

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lactate to pyruvate and pyruvate to acetyl-CoA; after diffusion through the cytoplasmic membrane, the H2 would be oxidized in the periplasm as described for H2 as electron donor. An involvement of periplasmic [Fe] hydrogenase in growth of Desulfovibrio vulgaris (Hildenborough) on lactate was also suggested by van den Berg et al. (1991). Reduction of the amount of this hydrogenase by means of antisense RNA resulted in a pronounced reduction of growth yields on lactate. The [NiFeSe] hydrogenase, which is located at the cytoplasmic aspect of the cytoplasmic membrane, might function as the H2-evolving hydrogenase, and the [Fe] and the [NiFe] hydrogenases are thought to function as the H2-oxidizing enzymes (Rohde et al., 1990). On the other hand, there are also arguments against free H2 as an obligatory intermediate in the catabolism of lactate. Important in this respect is the lack of a strong inhibition of lactate oxidation by an H2 atmosphere, unlike what should be expected for thermodynamic reasons in the H2-cycling model (e.g., Pankhania et al., 1986); furthermore, a Desulfovibrio mutant was isolated that does not grow on H2 plus sulfate but does grow on lactate plus sulfate (Odom and Wall, 1987). Also, there are other sulfate reducers growing well on lactate and other substrates without possessing hydrogenase, e.g., Desulfobotulus sapovorans or Desulfococcus multivorans. Hydrogen production linked to the oxidation of lactate to pyruvate has been even shown to be an energy-dependent process (Pankhania et al., 1988). The investment of energy makes H2 cycling as a mode of energy conservation on lactate unlikely. It is true that, with pyruvate as growth substrate for Desulfovibrio vulgaris, H2 cycling was directly demonstrated by employing membrane-inlet mass spectrometry (Peck et al., 1987). However, under natural conditions, cycling by Desulfovibrio of H2 from pyruvate oxidation is probably not a significant reaction. In natural habitats, pyruvate is probably not a major free product of fermentative bacteria and a less important substrate, if at all, for sulfate reducers than lactate. With pyruvate added to artificial media, a rapid pyruvate:ferrodoxin oxidoreductase (PFOR) reaction may cause a burst of H2 which is then scavenged mainly by periplasmic hydrogenase (Tsuji and Yagi, 1980). To our understanding, the production of some H2 during growth on lactate and sulfate (Lupton et al., 1984) is neither a proof for H2 cycling nor a proof for a specific mechanism that controls the redox state of electron carriers involved in lactate oxidation. One possible explanation is that part of the reducing power during growth on lactate and sulfate simply diffuses off via a constitutive hydrogenase; the H2 partial pressure may reflect the degree of

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imbalance between electron-producing and electron-consuming reactions. Another explanation can be given in view of the capacity of Desulfovibrio species to grow by interspecies H2transfer in sulfate-free cocultures with methanogens. Lactate conversion to acetate, H2 and CO2 in the absence of sulfate seems to be one of the ecological roles of Desulfovibrio species (Bryant et al., 1977; Zellner et al., 1987; Zellner and Winter, 1987; section Fermentative growth and syntrophy). In the presence of sulfate, the H2evolving system (Pankhania et al., 1988) may not be completely suppressed and lead to a minor loss of reducing power as H2. Ethanol and Acetaldehyde Ethanol is a very common electron donor and carbon source for incompletely and completely oxidizing sulfate reducers (Table 1). Ethanol is oxidized via acetaldehyde to acetate, which may be further oxidized. Some Desulfovibrio species can oxidize choline to acetate and trimethylamine. There is some evidence that acetaldehyde formed as the first intermediate from choline degradation is oxidized to acetate via acetyl-CoA (Hayward, 1960). With primary alcohols as electron donors, some sulfate-reducing bacteria form strong smelling byproducts which might be chemical adducts of sulfide and aldehydes that are formed as free intermediates (F. Widdel, unpublished observation). During growth on ethanol plus sulfate, Desulfovibrio gigas and three other examined Desulfovibrio strains contained high NAD+dependent alcohol dehydrogenase activities. In lactate-grown cells, these activities were lower or absent. NAD+-dependent alcohol dehydrogenases have been purified from Desulfovibrio gigas and from Desulfovibrio strain HDv; the latter organism is now known as Desulfovibrio burkinensis (Ouattara et al., 1999). Both enzymes were oxygen-labile; the proteins were decameric, with subunits of 43 and 48 kDa, respectively, and contained zinc (Hensgens et al., 1993; Hensgens et al., 1995a). The first 21 Nterminal amino acids of the enzyme from Desulfovibrio strain HDv were identical to those of the alcohol dehydrogenase from Desulfovibrio gigas; on the basis of the N-terminal amino acid sequences, the enzymes are members of the socalled family III of alcohol dehydrogenases which is not related to the family that includes the major yeast and mammalian alcohol dehydrogenases (Reid and Fewson, 1994). The alcohol dehydrogenases from Desulfovibrio are only highly active toward short primary alcohols; unlike the decameric family III enzyme from

CHAPTER 1.22

Bacillus methanolicus, however, they show no activity with methanol. A molybdenum iron-sulfur protein from D. gigas was shown to have some aldehyde dehydrogenase activity (Turner et al., 1987). Somewhat later, aldehyde oxidation was studied in more detail (Kremer et al., 1988b). Coenzyme A or phosphate dependency was not found, indicating that acetyl-CoA and acetyl phosphate are not intermediates in the conversion of acetaldehyde to acetate (Kremer et al., 1988b). Furthermore, it was shown that acetaldehyde can be oxidized in Desulfovibrio gigas by two completely different enzymes, a molybdenum-containing enzyme which can be assayed with DCPIP as artificial electron acceptor, and a tungsten-containing enzyme reacting with benzylviologen as an acceptor; the latter is strongly stimulated by K+ ions (Kremer et al., 1988b). The synthesis of the enzymes appeared to be strongly affected by the presence of molybdate and tungstate in the growth media (Hensgens et al., 1994). Extracts of cells grown in the presence of both tungstate and molybdate have only very low levels of the DCPIP-dependent enzyme. The benzylviologenlinked tungsten-containing aldehyde dehydrogenase allows much faster growth with ethanol than the molybdenum enzyme. During growth on ethanol of Desulfovibrio gigas, in media without tungstate, transient excretion of acetaldehyde was observed. The molydenum-containing aldehyde oxidoreductase is a homodimer of subunits with 907 amino acid residues and contains a molybdopterin cofactor and two different [2Fe2S] centers. It is a member of the xanthine oxidase family, and it is the first molybdenum enzyme with a molybdopterin cofactor (the crystal structure of which was determined; Romão et al., 1995). The tungsten-containing aldehyde oxidoreductase of Desulfovibrio gigas was active towards several aldehydes. This enzyme consists of two subunits of 62 kDa and was found to contain approximately 0.7 W, 4.8 Fe and 3.2 labile S per subunit; EPR studies indicated the presence of a [4Fe-4S] center (Hensgens et al., 1995b). Most likely, the tungsten-containing aldehyde oxidoreductase is related to similar enzymes from hyperthermophilic archaea and from Gram-positive anaerobic bacteria (Kletzin and Adams, 1996; Romão et al., 1997; Hu et al., 1999). The presence of a tungsten-containing aldehyde dehydrogenase, supposedly using flavins as natural cofactors, in Desulfovibrio simplex was demonstrated in experiments with cell-free extracts (Zellner and Jargon, 1997). Other Monovalent Alcohols and Polyols Methanol is a less common electron donor for sulfate-

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

reducing bacteria and growth on methanol is usually slower. Enrichment cultures with methanol usually select for methanogenic bacteria, despite the presence of sulfate. Methanol can be used by some Desulfotomaculum species such as the mesophilic Desulfotosporosinus orientis (Klemps et al., 1985), the thermophilic Desulfotomaculum kuznetsovii (Nazina et al., 1988), a few Desulfovibrio species (e.g., Desulfovibrio carbinolicus; Nanninga and Gottschal, 1987), Desulfobacterium anilini (Schnell et al., 1989) and Desulfobacterium catecholicum (Szewzyk and Pfennig, 1987). The mechanism of methanol oxidation is unknown. Primary alcohols higher than ethanol, for instance 1-propanol and 1-butanol, can also act as H2 donors for sulfate-reducing bacteria. Oxidation by Desulfovibrio species is incomplete and leads to the formation of the corresponding acids (propionate, butyrate, respectively; Mechalas and Rittenberg, 1960). Desulfobulbus strains oxidize 1-propanol incompletely to acetate (Widdel and Pfennig, 1982). Species of other genera may oxidize these alcohols completely. Certain Desulfovibrio strains were shown to dehydrogenate a secondary alcohol such as 2propanol to acetone (Widdel, 1986; Zellner et al., 1989a; Tanaka, 1992) or 2-butanol to 2-butanone (Tanaka, 1992). Desulfococcus biacutus (Platen et al., 1990) and Desulfococcus multivorans strains except for the type strain (Widdel, 1988) oxidize 2-propanol completely to CO2. Metabolism of diols by Desulfovibrio strains involves either an initial oxidation of the primary alcohol group yielding an hydroxyaldehyde, or the dehydration of the diol to an aldehyde. Thus, 1,2-propanediol can be metabolized to acetate with lactaldehyde as a presumed intermediate, or to propionate via propanal (see Hansen, 1994). Oxidation of 1,3-propanediol leads to 3hydroxypropionate or to acetate production as the major product (Nanninga and Gottschal, 1987; Qatibi et al., 1991; Tanaka, 1990; Tanaka, 1992); oxidation of 1,4-butanediol and 1,5pentanediol yielded the corresponding hydroxyacids (Tanaka, 1992). Oppenberg and Schink (1990) suggested a pathway involving malonylsemialdehyde for the conversion of 1,3propanediol to acetate by Desulfovibrio strain OttPd1. Some Desulfovibrio species were shown to grow on glycerol (e.g., Stams et al., 1985; Kremer and Hansen, 1987; Nanninga and Gottschal, 1987; Ollivier et al., 1988). In two marine Desulfovibrio strains, glycerol is degraded to acetate and CO2 via glycerol-3-phosphate, dihydroxyacetone phosphate and subsequent reactions known from the glycolytic pathway (Kremer

709

and Hansen, 1987). Desulfovibrio carbinolicus oxidizes glycerol to 3-hydroxypropionate (Nanninga and Gottschal, 1987). In the case of Desulfovibrio fructosovorans, acetate is the normal product during sulfate reduction, but during syntrophic growth with a methanogenic archaeon as H2 scavenger, glycerol is oxidized to 3-hydroxypropionate (Qatibi et al., 1998). Sugars Batch enrichment cultures with sugars commonly select for fermentative bacteria rather than for sulfate reducers, due to faster growth of the former. Nevertheless, some species of sulfate reducers isolated on other substrates were shown to use fructose in the absence or presence of sulfate (Klemps et al., 1985; Ollivier et al., 1988; Zellner et al., 1989a; Trinkerl et al., 1990). Desulfotomaculum nigrificans was originally reported to utilize glucose (Campbell et al., 1957; Akagi and Jackson, 1985). However, later growth tests with filter-sterilized sugars revealed that fructose rather than glucose is utilized by this species. When glucose had been autoclaved rather than filter sterilized, growth was observed, indicating partial conversion to a utilizable sugar, probably fructose (Klemps et al., 1985). Acetone Desulfococcus biacutus (Platen et al., 1990) and Desulfococcus multivorans strains other than the type strain (Widdel, 1988) used acetone that was completely oxidized to CO2. Desulfobacterium cetonicum is the other known sulfate-reducing bacterium that can grow with acetone as sole source of carbon and energy (Galushko and Rozanova, 1991). Acetone degradation was shown to depend on CO2 in cell suspensions of Desulfococcus biactus. Enzyme studies with the same microorganism indicated that it metabolized acetone via initial carboxylation to acetoacetyl-CoA. The latter is then thiolytically cleaved to two acetyl-CoA, which are further oxidized to CO2. The energy gain with acetone as substrate is low because degradation requires carboxylation and activation to acetoacetyl-CoA (Platen et al., 1990; Janssen and Schink, 1995a). Similar results were obtained for acetone metabolism of Desulfobacterium cetonicum (Janssen and Schink, 1995b). An ATP-dependent carboxylation of acetone under anaerobic conditions was also demonstrated in cell-free extracts of the photosynthetic bacterium Rhodobacter capsulatus (Birks and Kelly, 1997) and other bacteria (Ensign et al., 1998). Recently, an enrichment culture of sulfatereducing bacteria was described that could utilize the long-chain ketones hexadecan-2-one and 6,10,14-trimethylpentadecan-2-one. The oxida-

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tion of these ketones also involved a carboxylation reaction (Hirschler et al., 1998). Glycolate Glycolate is a widespread byproduct of autotrophic organisms (e.g. cyanobacteria and algae) in oxic environments with limiting CO2 concentrations. Ribulose-1,5-bisphosphate carboxylase of the Calvin cycle may incorporate O2 instead of CO2 in the substrate, such that one moiety is released as glycolate. It can be oxidized completely to CO2 by Desulfofustis glycolicus, an organism which was isolated from marine sediment (Friedrich et al., 1996). With methylene blue as an electron acceptor, a rather high activity of a membrane-bound glycolate dehydrogenase was detected (Friedrich and Schink, 1995). Malate, Fumarate, Succinate and Other Dicarboxylic Acids Dicarboxylic acids known from the citric acid cycle are relatively common substrates of incompletely and completely oxidizing sulfate reducers (Postgate, 1984a; Postgate, 1984b; Widdel, 1988). Growth on fumarate and malate is usually faster than on succinate. Some species may utilize only one or two of these compounds because certain transport systems might be limited or lacking. Growth yields of Desulfovibrio strains on succinate are far lower than on malate (Kremer et al., 1989). This may be explained by a partial investment of the conserved energy for reverse electron transport from the oxidation of succinate (fumarate/ succinate, E0¢ = +0.033 V). In various Desulfovibrio strains, the C4dicarboxylic acids are oxidized via a reaction sequence with an NADP+-dependent malic enzyme (a decarboxylating malate dehydrogenase); the activity was dependent on divalent cations (Mn2+ or Mg2+) and stimulated by K+ (Kremer et al., 1989). The NADP+-dependent malic enzyme of Desulfovibrio gigas was found to be a monomeric 45-kDa protein (Chen et al., 1995). The C5 and C7 dicarboxylic acids, glutarate and pimelate, respectively, can serve as substrates for some complete oxidizers (Bak and Widdel, 1986b; Imhoff-Stuckle and Pfennig, 1983; Schnell et al., 1989; Szewzyk and Pfennig, 1987). Amino Acids Utilization of amino acids as electron donors and carbon sources mainly has been reported for marine species. They include several Desulfovibrio strains; alanine utilization seems to be widespread and has been reported for Desulfotomaculum ruminis (Coleman, 1960), Desulfovibrio salexigens (Zellner et al., 1989a; van Niel et al., 1996), Desulfovibrio strains 20020

CHAPTER 1.22

and 20028 (Stams et al., 1985), Desulfovibrio acrylicus (van der Maarel et al., 1996c), and Desulfovibrio zosterae (Nielsen et al., 1999). Desulfocella halophila, which was isolated from sediment of the Great Salt Lake, is also able to use L-alanine as an electron donor (Brandt et al., 1999). Desulfovibrio acrylicus and Desulfovibrio strains 20020 and 20028 also have been shown to utilize serine, glycine and cysteine; even other amino acids are used by Desulfovibrio strains 20020 and 20028. Several Desulfobacterium strains use glutamate (ImhoffStuckle and Pfennig, 1983; Bak and Widdel, 1986b; Brysch et al., 1987; Szewzyk and Pfennig, 1987; Heijthuijsen and Hansen, 1989; Schnell et al., 1989; van der Maarel et al., 1996a; Rees et al., 1998). Some other amino acids were also utilized by Desulfobacterium strain PM4, the Desulfobacterium-like strain WN, and Desulfobacetrium vacuolatum (Heijthuijsen and Hansen, 1989; van der Maarel et al., 1996a; Rees et al., 1998). Desulfovibrio aminophilus, which was isolated from an anaerobic lagoon of a dairy wastewater plant, degraded six amino acids including alanine in the presence of sulfate (Baena et al., 1998). L-Alanine was found to be oxidized to pyruvate by an NAD+-dependent alanine dehydrogenase in Desulfovibrio strains 20020 and 20028 and in Desulfotomaculum ruminis (Stams and Hansen, 1986). Furfural Desulfovibrio furfuralis has been isolated with furfural; several other, previously known species of this genus also turned out to use this compound (Folkerts et al., 1989). On the basis of feeding experiments with 13 C-labeled furfural, a reaction sequence was postulated for the breakdown of the substrate in D. furfuralis with succinic semialdehyde as a key intermediate (Folkerts et al., 1989); furfuryl alcohol and 2-furoic acid transiently accumulated in the culture supernatants as important intermediates. Methylated N- and S-compounds (Glycine, Betaine, Dimethylsulfoniopropionate and Dimethylsulfide) Glycine betaine (trimethylglycine) is widely used as an osmolyte in many bacteria (for summary, see Galinski, 1995). Dimethylsulfoniopropionate is an osmolyte in many marine algae. Growth with glycine betaine as organic substrate has been demonstrated for a number of isolates belonging to the Desulfobacterium/ Desulfobacter cluster of the d-Proteobacteria; they include Desulfobacterium autotrophicum, Desulfobacterium niacini, Desulfobacterium vacuolatum, a strain named WN which clusters in

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

between Desulfobacterium and Desulfobacter (Heijthuijsen and Hansen, 1989; van der Maarel et al., 1996a), and Desulfospira joergensenii (Finster et al., 1997a). Glycine betaine (trimethylglycine) was demethylated to dimethylglycine as end product. It was speculated that the oxidation of the methyl group in these strains, which contain CO dehydrogenase, is mediated via the methyl branch of the oxidative C1-pathway normally used for the oxidation of the methyl moiety of the acetyl group in acetyl-CoA (Heijthuijsen and Hansen, 1989). Most organisms that can oxidize glycine betaine also grow by demethylation of dimethylsulfoniopropionate (DMSP) with 3-methylthiopropionate as the product; Desulfobacterium autotrophicum did not grow on DMSP (van der Maarel et al., 1996a). In cell extracts of DMSP-grown strain WN and other strains, a high DMSP:tetrahydrofolate methyltransferase activity was detected (Jansen and Hansen, 1998). Certain sulfatereducing bacteria have been shown to metabolize DMSP in a different way, namely by cleaving the DMSP to acrylic acid and dimethylsulfide and by reducing the acrylate to propionate (van der Maarel et al., 1996c; see below for a discussion of acrylate reduction). Dimethylsulfide is a widespread degradation product of dimethylsulfoniopropionate in marine environments. As a trace gas in the atmosphere, dimethylsulfide leads to sulfuric acid that forms condensation nuclei for water and thus influences cloud formation. Dimethylsulfide oxidation by mesophilic sulfate-reducing bacteria from marine sediments was inferred from experiments with labeled substrates and inhibitor studies (Kiene et al., 1986). Oxidation of dimethylsulfide by a thermophilic Desulfotomaculum strain has been reported (Tanimoto and Bak, 1994) but utilization of dimethylsulfide by pure cultures of mesophilic sulfate reducers remains to be demonstrated. Polar Aromatic Compounds (Non-Hydrocarbons) The utilization of various nonfermentable aromatic compounds in the absence of O2 or nitrate seems to be one of the domains of sulfatereducing bacteria. In contrast, aromatic compounds with more than two hydroxyl groups (e.g. gallic acid, pyrogallol or phloroglucinol) are readily degraded by fermentative bacteria (Schink and Pfennig, 1982; Schink, 1988a; Schink, 1988b). Several new types of sulfatereducing bacteria have been directly isolated with aromatic compounds (Widdel, 1980; Imhoff-Stuckle and Pfennig, 1983; Widdel et al., 1983; Bak and Widdel, 1987; Szewzyk and Pfennig, 1987; Schnell et al., 1989; Kuever et al., 1993; Gorny and Schink, 1994). Most of these

711

isolates are very versatile sulfate reducers that also use many aliphatic compounds. Benzoate is the most commonly and most readily utilized aromatic substrate. Representatives of other classes of aromatic compounds oxidized by sulfate reducers are phenol, p-cresol (Bak and Widdel, 1986b), aniline (Schnell et al., 1989), and the N-heterocyclic compounds nicotinate (Imhoff-Stuckle and Pfennig, 1983), indole and quinoline (Bak and Widdel, 1986a). An overview of non-hydrocarbon aromatic substrates utilized by pure cultures of sulfate-reducing bacteria is presented in Table 5. So far, most sulfate-reducing bacteria that degrade aromatic compounds are complete oxidizers. The only known exception is Desulfovibrio inopinatus, which degrades the relatively oxidized compound hydroxyhydroquinone (1,2,4-trihydroxybenzene) incompletely to acetate (Reichenbecher and Schink, 1997). Little is known about reactions at the aromatic ring in sulfate-reducing bacteria. Aerobic bacteria employ oxygenases which require O2 (as cosubstrate) to activate (hydroxylate) and cleave the aromatic ring. The pathways in the anaerobic sulfate-reducing bacteria are therefore expected to be completely different from those of aerobic bacteria and to involve novel biochemical reactions. Most insights into the degradative pathways of aromatic compounds under anoxic conditions were obtained from studies with denitrifying and phototrophic bacteria (for overview see e.g., Berry et al., 1987; Evans and Fuchs, 1988; Tschech, 1989a; Heider and Fuchs, 1997a; Heider and Fuchs, 1997b; Harwood et al., 1999). Important principles of aromatic compound degradation recognized in the nonsulfatereducing bacteria are that the degradative pathways can be classified into three categories. First, many aromatic compounds are converted via so-called peripheral reactions to a central intermediate, benzoyl-CoA. Second, a central sequence of reactions abolishes aromaticity of benzoyl-CoA and leads to ring cleavage. Third, certain polyhydroxybenzoates or polyhydroxybenzenes undergo reactions that lead to aliphatic intermediates without the involvement of benzoyl-CoA. Examples of peripheral reactions are phosphorylation/carboxylation to convert phenol to p-hydroxybenzoate (Knoll and Winter, 1989; Tschech and Fuchs, 1989b; Lack and Fuchs, 1992; Lack and Fuchs, 1994), the involvement of an aoxidation reaction in the conversion of phenylacetate to benzoyl-CoA (Mohamed et al., 1993), the reductive removal of hydroxyl groups (Tschech and Schink, 1986; Gibson et al., 1997). Free benzoate is simply activated to benzoylCoA in an ATP-consuming reaction (Geissler et al., 1988; Altenschmidt et al., 1991). Some

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R. Rabus, T.A. Hansen and F. Widdel

CHAPTER 1.22

Table 5. Sulfate-reducing bacteria with the capacitiy to use aromatic compounds as growth substrates. Organism Non-hydrocarbon aromatic compounds Desulfonema magnum Desulfococcus niacimb Desulfobacterium indolicumc Desulfobacterium phenolicumd Desulfobacterium catecholicume Desulfobacterium anilinic strain Cat2c strain SAXc Desulfotomaculum strain Grollf Desulfovibrio inopinatus Aromatic hydrocarbons “Desulfobacula toluolica” strain PRTOL1g strain mXyS1c strain oXyS1h strain NaphS2h

Aromatic substratea

Benzoate, 4-hydroxybenzoate, phenylacetate, 3-phenylpropionate, hippurate Nicotinic acid, 3-phenylpropionate Indole, 2-aminobenzoate, quinoline Phenol, p-cresol, benzoate, phenylacetate, indole, 4-hydroxyphenylacetate, 2-hydroxybenzoate, 4-hydroxybenzoate, phenylalanine, 2-aminobenzoate Catechol, resorcinol, 4-hydroxybenzoate, hydroquinone, benzoate 2-aminobenzoate, protocatechuate, phloroglucinol, pyrogallol Aniline, 2-aminobenzoate, 4-aminobenzoate, indolylacetate, quinoline Phenol, catechol, m-cresol, p-cresol, benzoate, phenylacetate, phenylpropionate 4-hydroxybenzoate, 3,4-dihydroxybenzoate, phenylalanine Benzoate, p-hydroxybenzoate, phenol, phenylacetate, phenylalanine Catechol, phenol, m-cresol,, p-cresol, benzoate, 3-hydroxybenzoate, benzaldehyde benzyl alcohol, phenylacetate, phenylpropionate Hydroxyhydroquinone (1,2,4-trihydroxybenzene) Toluene, p-cresol, benzaldehyde, benzoate, phenylacetate, p-hydroxybenzoate p-hydroxybenzaldehyde Toluene, p-cresol, benzaldehyde, benzoate, phenylacetate, phenylpropionate p-hydroxybenzoate Toluene, m-xylene, m-ethyltoluene, m-isopropyltoluene, benzoate, m-methylbenzoate Toluene, o-xylene, o-ethyltoluene, o-methylbenzyl alcohol benzoate, o-methylbenzoate, benzylsuccinate Naphthaline, 2-naphthoate, benzoate

Reference

Widdel etal., 1983 Imhoff-Stuckle and Pfennig, 1983 Bak and Widdel, 1986a Bak and Widdel, 1986b Szewzyk and Pfennig, 1987 Schnell et al., 1989 Schnell et al., 1989 Drzyzga et al., 1993 Kuever et al., 1993 Reichenbecher and Schink, 1997 Rabus et al., 1993 Beller et al., 1996 Harms et al., 1999 Harms et al., 1999 Galushko et al., 1999

a

Information about additional aromatic substrates is provided in the respective references. Has to be reclassified as “Desulfobacterium niacim” (J. Kuever, F. A. Rainey and F. Widdel, personal communication). c Has to be classified/reclassified as new genus (J. Kuever, F. A. Rainey and F. Widdel, personal communication). d Has to be reclassified as “Desulfobacula phenolicum” (J. Kuever, F. A. Rainey and F. Wuddel, personal communication). e Has to reclassified (J. Kuever, F. A. Rainey and F. Widdel, personal communication). f Has been classified as Desulfotomaculum gibsoniae (Kuever et al., 1999). g Has to be classified as a new species of the genus Desulforhabdus (J. Kuever, F. A. Rainey and F. Widdel, personal communication). h Has to be classfied as a new species of the genus Desulfosarcina (J. Kuever, F. A. Rainey and F. Widdel, personal communication). i Strain NaphS2 affiliates closely with strain mXyS1 (Galushko et al., 1999) and will therefore be classified into the same new genus (J. Kuever, F. A. Rainey and F. Widdel, personal communication). b

peripheral reactions for the degradation of aromatic compounds also have been suggested thus far in sulfate-reducing bacteria. Degradation of aniline by Desulfobacterium anilini is initiated by a carboxylation probably yielding 4aminobenzoate, which via ligation with acetylCoA and reductive deamination is supposed to yield benzoyl-CoA (Schnell and Schink, 1991). p-Cresol was first suggested to be converted to p-hydroxybenzyl alcohol by an anaerobic pcresol methylhydroxylase (McIntire et al., 1985;

Suflita et al., 1989); further oxidation to the corresponding aldehyde and acid, ligation with coenzyme A and reductive dehydroxylation (or vice versa) could yield benzoyl-CoA. Such a pathway would be in agreement with the ability of p-cresol-utilizing sulfate reducers to grow with benzoate. Furthermore, an anaerobic degradation of m-cresol by Desulfotomaculum strain (Groll) is proposed to proceed via a methylgroup oxidation to 3-hydroxybenzoate because the latter compound was detected in m-cresol-

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

degrading cultures (Londry et al., 1997). However, in the light of recent findings about the anaerobic activation of toluene by methyl condensation with fumarate (see Aromatic Hydrocarbons) methyl hydroxylation reactions may be questioned and reactions analogous to toluene activation may be assumed. Indeed, an activation of m-cresol by a fumarate-dependent reaction to 3-hydroxybenzylsuccinate was demonstrated in cell-free extracts of Desulfobacterium cetonicum (Müller et al., 1999). Catechol degradation by Desulfobacterium strain Cat2 was proposed to be initiated by a carboxylation to protocatechuate, because high activities of a protocatechuate decarboxylase and low activities of an ATP/ HCO3--dependent protocatechuyl-CoA-forming enzyme synthetase could be measured in extracts of catechol grown cells. Further degradation to benzoyl-CoA would involve reductive dehydroxylation reactions (Gorny and Schink, 1994). The further metabolism of benzoyl-CoA has been studied most intensely in denitrifying Thauera aromatica strain K172. The stable aromatic state is abolished by benzoyl-CoA reductase. This novel enzyme contains FAD as prosthetic group and uses ferredoxin as the natural electron donor. It requires two ATP to generate and transfer two electrons into the ring of benzoyl-CoA to yield cyclohexa-1,5-diene-1carboxyl-CoA (Boll and Fuchs, 1995; Boll and Fuchs, 1998); the first electron has to have an extremely negative redox potential. Different enzymes have been measured and isolated from Thauera aromatica (Laempe et al., 1998) and Rhodopseudomonas palustris (Perrotta and Harwood, 1994; Pelletier and Harwood, 1998) that are involved in further reduction and cleavage of the ring structure to yield the open chain pimelyl-CoA, which can be further degraded to acetyl-CoA via reactions such as b-oxidation. Thus, somewhat different pathways for benzoate degradation are employed by these two organisms (Harwood and Gibson, 1997; Harwood et al., 1999), suggesting that variations of pathways exist for the anaerobic degradation of benzoate. Considering the high energy requirement of anaerobic benzoate degradation in the aforementioned microorganisms, it appears rather unlikely that sulfate-reducing bacteria with their relatively low ATP yield employ the same reactions for ring reduction. Desulfococcus multivorans required selenite in addition to molybdate for the degradation of benzoate, but not for growth on aliphatic substrates (Widdel, 1980). However, neither the role of these trace elements in Desulfococcus nor the pathway of benzoate degradation is known. Heterocyclic aromatic compounds utilized by sulfate-reducing bacteria are nicotinate (Imhoff-

713

Stuckle and Pfennig, 1983), indole and quinoline (Bak and Widdel, 1986a). Desulfobacterium niacini requires traces of selenite for the oxidation of nicotinate (Imhoff-Stuckle and Pfennig, 1983), as Clostridium barkeri does for fermentation of this compound. In the latter, the degradation of nicotinic acid is initiated by a conversion to 6hydroxynicotinate via nicotinate dehydrogenase, which is probably a selenoenzyme (Imhoff and Andreesen, 1979). Nicotinate dehydrogenase, also detected in Desulfobacterium niacini (W. Buckel, personal communication), may explain the selenium requirement. Aromatic Hydrocarbons Aromatic hydrocarbons as apolar molecules are biochemically less reactive than their aromatic counterparts carrying functional groups. Degradation of aromatic hydrocarbons under anaerobic conditions was long considered to be impossible. However, studies with anaerobic sediment and enrichment cultures of mixed methanogenic cultures (GribcGalic and Vogel, 1987), denitrifying (Kuhn et al., 1988) and sulfate-reducing bacteria (Edwards et al., 1992) demonstrated that aromatic hydrocarbons such as toluene were indeed degradable under anoxic conditions. The first pure cultures that could anaerobically degrade toluene were obtained under denitrifying (Dolfing et al., 1990; Altenschmidt and Fuchs, 1991; Evans et al., 1991; Schocher et al., 1991) and ferric-iron reducing (Lovley et al., 1990) conditions. The first pure culture of a toluene-degrading sulfate-reducing bacterium, “Desulfobacula toluolica,” was isolated from marine sediment (Rabus et al., 1993). This new isolate oxidized toluene completely to CO2 according to equation (16), as demonstrated by measurement of the degradation balance. Another toluene-degrading sulfate reducer, strain PRTOL1, was isolated from fuelcontaminated subsurface soil (Beller et al., 1996). 2-

C 6 H5 CH3 + 4.5SO4 + 2H + + 3H 2 O Æ 7HCO3 + 4.5H 2 S o DG ¢ = -205 kJ mol toluene

(16)

A marine enrichment culture that grew anaerobically on crude oil with concomitant sulfate reduction to sulfide (Rueter et al., 1994) was the source for the isolation of the o-xylenedegrading strain oXyS1 and the m-xylenedegrading strain mXyS1 (Harms et al., 1999). Both strains also used toluene for growth by sulfate reduction. Furthermore, strain oXyS1 oxidized o-ethyltoluene, and strain mXyS1 oxidized m-ethyltoluene and m-isopropyltoluene anaerobically. Sulfate-reducing strain NaphS2 was isolated as the first pure culture which can

714

R. Rabus, T.A. Hansen and F. Widdel

CHAPTER 1.22

COO– COO– CH3

COO– Fumarate

COSCoA COO–

Toluene

Benzylsuccinate

utilize the bicyclic aromatic hydrocarbon naphthalene (Galushko et al., 1999). Anaerobic degradation of other aromatic hydrocarbons with sulfate as electron acceptor has been demonstrated in enriched sediment communities, but not so far in pure cultures. These hydrocarbons are benzene (Lovley and Phillips, 1995b; Phelps et al., 1996) and the polyaromatic hydrocarbons phenanthrene and fluorene (Coates et al., 1997). Oxidation was shown by the formation of 14CO2 from the 14C-labeled hydrocarbon substrates. The best known anaerobic pathway of an aromatic hydrocarbon is that of toluene. Understanding of anaerobic toluene metabolism has greatly benefited from studies with denitrifying bacteria. Benzylsuccinate, first identified as a metabolite in toluene-grown cultures of a denitrifyer (Evans et al., 1992), a sulfate-reducing enrichment culture (Beller et al., 1992) and Desulfobacula toluolica (Rabus and Widdel, 1995), was shown to be the initial activation product in denitrifying bacteria (Biegert et al., 1996; Beller and Spormann, 1997a). It was formed from toluene and fumarate. Fumaratedependent formation of benzylsuccinate from toluene was subsequently reported with permeabilized cells of sulfate-reducing strain PRTOL1 (Beller and Spormann, 1997b) and in cell-free extracts of Desulfobacula toluolica (Rabus and Heider, 1998). The further demonstration of benzylsuccinate formation in a toluene-utilizing phototroph that is unrelated to denitrifying or sulfate-reducing bacteria (Zengler et al., 1999b) suggests that this is a general anaerobic activation mechanism for toluene, a naturally widespread trace hydrocarbon (Heider et al., 1999). Genetic analysis of genes underlying the benzylsuccinate-forming enzyme (benzylsuccinate synthase) indicates that this is a glycyl radical enzyme (Coschigano et al., 1998; Leuthner et al., 1998); the radical is supposed to attack the methyl group of toluene yielding a benzyl radical which then combines with fumarate (Fig. 16). Further degradation of benzylsuccinate is proposed to proceed via reactions analogous to b-oxidation of a-methyl-

Fig. 16. Anaerobic, fumaratedependent activation of toluene to benzylsuccinate in Desulfobacula toluolica and strain PRTOL1. Further degradation of benzylsuccinate to the central intermediate benzoylCoA is not completely understood.

Benzoyl-CoA

branched fatty acids and to yield benzoyl-CoA as a central intermediate. In agreement with this, toluene-utilizing sulfate-reducing bacteria can also grow on benzoate. Hints as to the initial reaction in anaerobic degradation of naphthalene were obtained from enriched sediment communities under sulfatereducing conditions. The finding of 2-naphthoate (naphthalene-2-carboxylate) suggested a carboxylation as the initial activation of the bicyclic aromatic hydrocarbon (Zhang and Young, 1997). In agreement with this, naphthalenedegrading strain NaphS2 is able to grow on 2naphthoate, but not on 1-naphthoate (Galushko et al., 1999). A different initial mechanism of anaerobic napthalene degradation was suggested in a study of freshwater microcosms under conditions of sulfate reduction; in these communities, a naphthol (isomer unknown) was detected as a possible intermediate (Bedessem et al., 1997). Saturated Hydrocarbons Saturated hydrocarbons (n-alkanes, branched-chain alkanes and cycloalkanes) are the chemically least reactive organic compounds. The chemical recalcitrance is explained by the exclusive presence of apolar s bonds. Because of these structural properties and the fact that aerobic bacteria initiate alkane activation always with O2 as co-substrate (monoxygenase reaction), the possibility of an anaerobic alkane oxidation has often been doubted. Nevertheless, evidence for the anaerobic oxidation of alkanes in enriched microbial communities and pure cultures has been repeatedly provided. In the 1940s, enrichment cultures and pure cultures of Desulfovibrio strains were reported to grow or to reduce sulfate with long-chain alkanes (Novelli and ZoBell, 1944; Rosenfeld, 1947). The techniques available at that time to guarantee strictly anoxic conditions in the experiments were not described in detail. The cultures have not been preserved. In experiments with suspensions of other Desulfovibrio species, sulfate reduction was stimulated by octadecane and a

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

715

Table 6. Sulfate-reducing bacteria with the capacitiy to use aliphatic hydrocarbons as growth substrates. Aliphatic hydrocarbon utilizeda Organism

Optimum temperature (∞C)

n-Alkanes

1-Alkenes

Reference

Hxd3 Pnd3c TD3cd

28–30 30 55–65

C14, C16, C18 C14, C16, C18

AK01e

26–28

C12–C20 C14–C17 C6–C16, 3-methyloctane C13–C18

Aeckersberg et al., 1991 Aeckersberg et al., 1994, 1998 Rueter et al., 1994 Ehrenreich 1996 So and Young, 1999a

b

C15, C16

a

So far tested; more detailed information on growth substrates can be obtained from the respective references. Has tentatively been classified as “Desulfobacterium oleovorans” (Aeckersberg et al., 1991). The genus name has to be reclassified (J. Kuever, F. A. Rainey and F. Widdel, personal communication). c Has to be classified as a new genus (J. Kuever, F. A. Rainey and F. Widdel, personal communication). d Has been tentatively classified as “Desulfothermus naphthae” (Ehrenreich, 1996). e Has to be classified into the same new genus as strain Hxd3 (J. Kuever, F. A. Rainey and F. Widdel, personal communication). b

–30

∆G (kJ/mol SO42–)

small part (around 0.4%) of 14C-labeled alkane was recovered as CO2 (Davis and Yarbrough, 1966). The possibility of anaerobic alkane oxidation with sulfate was again examined in connection with a study of sulfate reducers in oil fields. A sulfate-reducing bacterium that nutritionally and morphologically differed from Desulfovibrio was isolated with hexadecane (Aeckersberg et al., 1991). Quantitative degradation experiments in anoxic, fused glass (air-excluded) ampullas showed that up to ca. 90% of the added hexadecane was oxidized with sulfate. A control experiment with a Desulfovibrio strain did not reveal alkane utilization. Three other pure cultures of alkane-degrading sulfate-reducing bacteria, two mesophilic strains (Aeckersberg et al., 1998; So and Young, 1999a) and a thermophilic strain (Rueter et al., 1994) were subsequently described. The range of alkanes utilized by these isolates and other characteristics are summarized in Table 6. Anaerobic utilization of various longchain n-alkanes was also observed with enriched communities in marine sediment under conditions of sulfate reduction (Caldwell et al., 1998). The biochemical problem in anaerobic alkane degradation is the first step, the activation of an apolar molecule, rather than in the free energy change of the overall reaction. With the exception of methane oxidation (see next section), the amount of free energy per mol of sulfate reduced with alkanes (see Fig. 17) is comparable to that available from acetate or propionate oxidation. The activation has to start with a cleavage of a C-H bond that is not activated. A resulting alkyl radical would not have the possibility for stabilization by delocalization, as in the case of an aryl radical (e.g., benzyl radical; see preceding section). Studies on changes in the cellular fatty acid composition in response to the growth substrate provided first hints at possible activation reactions of alkanes in a sulfate-reducing bacterium, strain Hxd3 (Aeckersberg et al., 1998). If cells

gaseous

liquid

–40

–50

–60

–70

2

4

6

8

10

12

14

16

C-Atoms per chain (n) Fig. 17. Free energy change of sulfate reduction with nalkanes of various chain lengths (methane through hexadecane) at 25∞C, pH = 7, SO42- and HCO3- concentrations = 10-2 M, and HS- concentration = 10-3 M. Individual stoichiometric equations are derived from CnHn+2 + (3n+1)/4 SO42Æ n HCO3- + (3n+1)/4 HS- + (n–1)/4 H+ + H2O, with n being the number of carbon atoms per chain. Free energy values were calculated from data given by Dean (1992); Thauer et al. (1977) and Zengler et al. (1999a).

were grown on hexadecane (C16H34), the chains of cellular fatty acids were mainly C-odd. Conversely, cells grown on heptadecane (C17H36) contained mainly C-even fatty acids in the lipid fraction. It was concluded from these results that the alkane chain was altered by a C1-unit during activation, possibly by the terminal addition of a C1-compound. However, with a second, phylogenetically related alkane-degrading sulfate reducer, strain Pnd3, a C-even alkane yielded C-even fatty acids, and a C-odd alkane yielded C-odd fatty acids. Assuming a mechanism principally as in strain Hxd3, activation at the second carbon atom was proposed as one possible explanation for the findings with strain Pnd3.

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R. Rabus, T.A. Hansen and F. Widdel

Also, addition to fumarate was discussed as hypothetical activation mechanism, which could differ from toluene activation by a lack of radical stabilization in the substrate molecule (Aeckersberg et al., 1998). Chemical analysis with a third alkane-degrading sulfate reducer, strain AK-01, yielded methyl-branched cellular fatty acids resulting from the n-alkane provided as substrate (So and Young, 1999b). Labeling studies suggested that a carbon compound, which is not derived from bicarbonate, is subterminally added to the alkane such that the terminal methyl group of the n-alkane becomes a methyl branch in the fatty acid formed via subsequent reactions. Methane Methane, the only existent stable C1-hydrocarbon, can be regarded as the first member of the homologous series of alkanes. It is chemically even somewhat more stable than higher alkanes. Methane is formed as an end product of anaerobic degradation processes involving methanogenic archaea in sediments that are depleted of electron acceptors other than CO2. Because of the important role of methane in the carbon cycle in aquatic habitats and on a global scale, the possibility of an anaerobic oxidation of this hydrocarbon has been frequently investigated. In anoxic marine habitats, sulfate would be the most important terminal electron acceptor for anaerobic methane oxidation. Hints on an anaerobic methane oxidation came mostly from biogeochemical investigations in marine sediments. Geochemical evidence is based on three different observations. First, methane in marine habitats often disappears far below the oxic zone, and the depth profile of the methane concentration exhibits a concaveup curvature, which indicates a methane sink (Devohl and Ahmed, 1981; Reeburgh, 1976; Barnes and Goldberg, 1976; Martens and Berner, 1977; Alperin and Reeburgh, 1984); an increase (“second maximum”) of the sulfate reduction rate in the depth profile was observed to coincide with the zone of anaerobic methane depletion (Alperin and Reeburgh, 1985; Iversen and Jørgensen, 1985; Reeburgh and Alperin, 1988; Hansen et al., 1998). Second, 13C/12C analyses are in favor of an anaerobic methane oxidation. Residual methane in the zone of its anaerobic depletion is 13C-enriched (and 2Henriched), indicating a biochemical consumption reaction (Alperin et al., 1988). In addition, inorganic carbon (CO2, HCO3-, CO32-) in the zone of methane depletion was shown to be relatively poor in 13C (Reeburgh, 1980; Reeburgh and Alperin, 1988; Blair and Aller, 1995); this finding

CHAPTER 1.22

suggested that oxidation of isotopically light methane added to the signature of the isotopically heavier background of inorganic carbon. Third, after addition of 14C-labeled methane to anoxic marine sediment cores or slurries, formation of radioactive CO2 could be measured (Reeburgh, 1980; Iversen and Blackburn, 1981; Alperin and Reeburgh, 1984; Alperin and Reeburgh, 1985; Iversen and Jørgensen, 1985; Hansen et al., 1998). The rates of anaerobic methane oxidation calculated from data of the biogeochemical investigations were always rather low; they ranged between 1 and 67 mmol · liter -1 · day -1, or were even lower. However, at a gas seep, volumetric sulfate reduction rates as high as 2.5 mmol · liter -1 · day -1 were attributed to methane as the electron donor (Aharon and Baoshun, 2000); this implies that methane oxidation at this site has the same rate. An organism that can consume methane anaerobically has not been enriched and isolated thus far. A partial conversion of 14CH4 to 14 CO2 during methanogenesis but no net oxidation of methane has been measured in cultures of methanogenic archaea (Zehnder and Brock, 1979; Zehnder and Brock, 1980), which provided the first hint of a “reverse methanogenesis”. Because biologically produced methane, which is usually used for labeling experiments, may contain traces of CO as a by-product, 14 C-methane was purified from this by-product and applied to active methanogenic bacteria (Harder, 1997). Again, a partial oxidation of methane without net consumption was demonstrated. The reaction was not detectable in cultures of sulfate-reducing and homoacetogenic bacteria. The assumption that anaerobic oxidation of methane is a reversed methanogenesis and catalyzed by methanogenic archaea (or at least by a phylogenetically closely related group) is supported by microbiological in situ analysis of bacterial populations on the basis of biomarkers and 16S rRNA gene sequences. Special isoprene lipids and hydrocarbons such as crocetane (2,6,11,15-tetramethylhexadecane) that occurred in the zone of methane depletion and exhibited an unusually low 13C/12C-ratio were assumed to belong to the methane-utilizing anaerobes (Elvert and Suess, 1999; Hinrichs et al., 1999); also retrieved 16S rRNA gene sequences forming a distinctive cluster within the Methanosarcinales were tentatively assigned to these microorganisms (Hinrichs et al., 1999). From these and earlier studies (Hoehler et al., 1994; Hansen et al., 1998), it was concluded that methane is not directly utilized by sulfatereducing bacteria, but rather by a group of archaea (eventually identified as methanogens)

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

that convert methane in a “reversed methanogenesis” to CO2 and an intermediate, possibly H2; the latter is then scavenged and kept at low concentration by the activity of sulfate-reducing bacteria. The free energy yield from anaerobic methane oxidation with sulfate near natural concentrations is relatively low (DG = –33 kJ/mol sulfate). This amount would have to be shared between two partners (Fig. 11). Assuming an equal share of the free energy with H2 as the intermediate (conditions see below following equations), the partial pressure of the latter would have to be around 0.12 Pa (corresponding to: 0.9 ·10 -9 M dissolved H2; E¢ = -0.269 V at pH 7.5) to render methane oxidation thermodynamically feasible. -

CH 4 + 3H 2 O Æ HCO3 + 4H 2 + H + DG = -15.7 kJ mol

(17)

2-

SO4 + 4H 2 + H + Æ HS - + 4H 2 O DG = -15.7 kJ mol 2-

(18)

-

Sum : CH 4 + SO4 Æ HCO3 + HS - + H 2 O (19) DG = -31.4 kJ mol (calculated for 25∞C; pH = 7.5; CH4 partial pressure = 105 Pa; H2 partial pressure = 0.12 Pa; SO42concentration = 2 ·10 -2 M; HCO3- concentration = 10-2 M; HS- concentration = 2 · 10 -3 M; activity coefficients of SO42-, HCO3- and HS- in seawater of 0.1, 0.5 and 0.5, respectively; data for calculation from Stumm and Morgan, 1981, and Thauer et al., 1989b). Measurement of H2 at partial pressures in the indicated range is technically possible. However, because the partial pressure is the result of a dynamic equilibrium between production and consumption, sampling procedures that affect substrate availability are expected to have a significant influence on the H2 partial pressure. Hydrogen partial pressures reported for conditions of sulfate reduction were 5 Pa in marine sediment (Sørensen et al., 1981), 0.17 Pa in sulfate-amended lake sediment (Lovley et al., 1982), and between 0.05 and 0.4 Pa in the anoxic seawater of Cariaco Trench (Scranton et al., 1984). Hence, the partial pressures determined in the latter samples would be roughly in the range required if anaerobic methane oxidation occurred via free H2. Sulfate reduction at the calculated very low H2 concentrations is expected to be very slow, even if sulfate-reducing bacteria are closely associated with the H2-producing partners. With the most favorable kinetic parameters reported for cells of sulfate-reducing bacteria, viz. a maximum rate (Vmax) of 90 mol H2 g-1 · h -1 (see Overview of

717

Principal Properties, Sulfate-Reducing Bacteria and Archaea in this Chapter) and a halfsaturation constant (KM) of 0.7 · 10 -6 mol ¥ H2 l-1 (Widdel, 1988), the specific rate (related to cell dry mass) of H2 oxidation would be 0.12 mol ¥ g-1 · h -1; hence the rate of sulfate reduction or methane oxidation would be 0.03 mol g-1 · h -1. (The rate at substrate concentrations KM is calculated by multiplication of the first-order rate constant, Vmax/KM, with the substrate concentration.) Since members of the Methanosarcinales are metabolically versatile, also a transfer of metabolites other than H2 may be assumed. However, organic compounds known as methanogenic substrates would require concentrations even lower than that of hydrogen to allow reverse methanogenesis and an approximately equal energy share of both partners (acetate, 3 · 10-11 M; concentrations of methanol and methylsulfide even lower). Hydrogen or electron carriers with midpoint potentials close to the redox potential calculated above (-0.269 V) would allow kinetically more favorable concentrations for a transfer of their oxidized and reduced forms between the partners. Special Inorganic Electron Donors (Other than H2) An economically important inorganic electron donor for sulfate-reducing bacteria is metallic iron. Oxidation of metallic iron with sulfate as electron acceptor is regarded as the principal reaction in anaerobic corrosion (Hamilton, 1985; Postgate, 1984c; Sequeira and Tiller, 1988; Von Wolzogen Kuhr and van der Vlught, 1934; Widdel, 1992). Anaerobic corrosion, a process with significant economic impact, has been frequently observed to cause pitting and destruction of pipelines and other iron and steel constructions exposed to sulfate-containing, oxygen-depleted waters. Because of the negative redox potential (Fe2+/Fe, E0 = -0.44 V; even more negative in carbonate-rich or sulfidic medium), iron can liberate H2 (2H+/H2, E0¢ = -0.41 V) in aqueous surroundings (according to 2 Fe + 2 H+ Æ 2 Fe2+ + H2) and may in this way indirectly act as an electron donor for sulfate-reducing bacteria that possess hydrogenase (Cord-Ruwisch and Widdel, 1986). However, a direct utilization of electrons (liberated according to Fe Æ Fe2+ + 2 e-) by cells associated with the iron surface and involving redox proteins at the cell surface (outer membrane) has been discussed as another mechanism in anaerobic corrosion (Van Ommen Kloeke et al., 1995; Widdel, 1992). Such a direct withdrawal of electrons may be kinetically more favorable than consumption of the electrochemically formed H2. Another inorganic, unique electron donor for dissimilatory sulfate reduction is phosphite

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R. Rabus, T.A. Hansen and F. Widdel

(H2PO3-), that has been used for the enrichment and isolation of a novel type of sulfate-reducing bacterium (Schink and Friedrich, 2000). The isolate is phylogenetically related to Desulfobacula and Desulfospira. Phosphite was oxidized to phosphate (H2PO4-). The natural role of this capacity is unknown. The occurrence of a “dissimilatory phosphate reduction” in natural habitats as a source of reduced phosphorous compounds is very unlikely, because the redox potentials of the reduction steps of phosphorus (ranging from +V to -III) are extremely low (E0¢ lower than -0.48 V; Schink and Friedrich, 2000; Widdel, 1992). Reduction of H+ to H2 would be easier to achieve (see redox potential above). Fermentative and Syntrophic Growth in the Absence of Sulfate In the absence of sulfate or other inorganic electron acceptors, several types of sulfate reducers can grow by fermentation of several organic substrates. Some Desulfovibrio species (for overview see Widdel, 1988), Desulfobacterium species (Brysch et al., 1987) and Desulfosarcina variabilis (Widdel, 1980) ferment fumarate and, with the exception of the latter species, malate; the fermentation products are succinate, acetate, CO2, and sometimes propionate. Pyruvate is easily fermented by many sulfatereducing bacteria that can use lactate. Pyruvate fermentation by Desulfovibrio desulfuricans produces acetate, CO2 and H2 (e.g., Postgate, 1984a; Stams et al., 1985). In Desulfovibrio sapovorans, which also ferments pyruvate but does not possess a hydrogenase (Widdel, 1980), lactate, acetate and CO2 are the expected fermentation products. Lactate or ethanol plus CO2 allow fermentative growth of some Desulfobulbus strains that form propionate and acetate (Laanbroek et al., 1982; Widdel and Pfennig, 1982). Propionate is formed in Desulfobulbus via a randomizing pathway involving a methylmalonyl-CoA:pyruvate transcarboxylase and free succinate as a symmetric molecule (Stams et al., 1984). This pathway is very similar to the one used by Propionibacterium except that the activation of succinate to succinyl-CoA is not directly linked to the formation of propionate from propionylCoA. The succinate pathway in the inverse direction also is used for the oxidation of propionate to acetate and CO2 in the presence of sulfate (see Propionate in this Chapter; Kremer and Hansen, 1988a; Fig. 15). Desulfovibrio desulfuricans can ferment choline to trimethylamine, ethanol, and acetate (e.g., Fiebig and Gottschalk, 1983). Desulfovibrio fructosovorans and Desulfotomaculum nigrificans fermented fructose in the absence of sulfate

CHAPTER 1.22

(Klemps et al., 1985; Ollivier et al., 1988). The former was shown to form succinate, acetate and ethanol. Furthermore, a fermentation of cysteine with liberation of sulfide and ammonia has been reported for a sulfate reducer, probably a Desulfovibrio strain (Senez and Leroux-Gilleron, 1954b). Desulfotosporosinus orientis grew slowly by converting formate, methanol, or the methyl groups of 3,4,5-trimethoxybenzoate via a homoacetogenic metabolism to acetate (Klemps et al., 1985). Lactate was fermented by this species to acetate as the only organic product, which is in agreement with the observed de novo acetate formation as it occurs in homoacetogenic bacteria. Also in Desulfobacterium species, fermentation of lactate and malate yielded an acetate to substrate ratio that can only be explained by an additional de novo synthesis of acetate from reducing equivalents and CO2 (Brysch et al, 1987; F. Widdel, unpublished observation). Desulfovibrio carbinolicus and Desulfovibrio fructosovorans ferment glycerol to 1,3-propanediol and 3-hydroxypropionate (Nanninga and Gottschal, 1987; Ollivier et al., 1988). The marine sulfate-reducing bacterium Desulforhopalus singaporensis was isolated from an anaerobic enrichment culture with taurine (2-aminoethanesulfonate) as the only source of carbon, energy, and nitrogen (Lie et al., 1999). The degradation of taurine, that includes a reduction of the oxidized sulfur, could be described by the following equation (22): -

2 + H3 N-CH 2 CH 2 CH 2 SO3 Æ CH3 COO+ + 2CO2 + 2 NH 4 + 2HS - + H +

(22)

Another sulfonate that was reported to be fermented by a sulfate reducer is cysteate (Laue et al., 1997a). The fermentation of cysteate by Desulfovibrio strain GRZCYSA could be approximated by the following equation (23):

(

+

)

2 -O3 S -CH 2 CH NH3 COO- + 2H 2 O Æ 2CH3 COO- + 2CO2 + SO4 + + HS - + 2 NH 4 + H +

2-

(23)

Desulfovibrio species may grow with ethanol or lactate in the absence of sulfate if co-cultured with H2-scavenging methanogenic bacteria (Bryant et al., 1977). In these syntrophic associations, the sulfate reducers serve as syntrophic, H2-producing acetogenic bacteria. Without a H2scavenging partner in the absence of sulfate,

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

Desulfovibrio forms H2-partial pressures of up to 1.5 kPa, without growth (Pankhania et al., 1988). Hydrogen formation in Desulfovibrio from lactate in the absence of sulfate was inhibited by protonophores and inhibitors of protontranslocating ATPase, whereas H2 formation from pyruvate was not inhibited under such conditions (Pankhania et al., 1988). This observation indicated that the reducing equivalents from lactate dehydrogenation were converted to H2 in an energy-driven process, as also suggested by the concerning redox couples (pyruvate/lactate, E0¢ = –0.190 V; 2 H+/H2, E0¢ = –0.414 V). This process may be a reversed electron transport driven by the proton gradient, as in substrate oxidation of other syntrophic bacteria (Schink, 1997). The assumption of a chemiosmotically driven dehydrogenation of lactate is further supported by the fact that lactate dehydrogenase is associated with the cytoplasmic membrane (see section on lactate). The energy for lactate dehydrogenation in Desulfovibrio is presumably derived from the subsequent exergonic conversion of pyruvate via acetyl-CoA (Acetyl-CoA + CO2/pyruvate, E0¢ = –0.5 V) and acetyl phosphate to free acetate; the latter step allows substratelevel phosphorylation and generation of a proton gradient through ATP hydrolysis at the ATPase. In a coculture with a methanogenic bacterium, a Desulfovibrio species converted choline to trimethylamine, acetate, and H2; the latter was used by the methanogenic partner (Fiebig and Gottschalk, 1983). Desulfonema limicola, Desulfosarcina variabilis, and species of the genera Desulfobulbus, Desulfobacterium, and Desulfotomaculum did not grow in cocultures with methanogens in sulfate-free medium on lactate, ethanol, fatty acids or benzoate, even though the tested sulfate reducers possessed hydrogenase (C. Schneider and F. Widdel, unpublished observation). Apparently, there is no special mechanism in these sulfate reducers for the transfer of reducing equivalents to the redox-level of H2, as it probably occurs in Desulfovibrio (Pankhania et al., 1988). However, further sulfate-reducing bacteria that grow syntrophically with H2 scavengers were detected during investigations on methanogenesis from propionate. The propionate-utilizing, syntrophic partners, that were named as Syntrophobacter species, turned out to be sulfate-reducing bacteria and members of the d-subclass (Dörner, 1992; Harmsen et al., 1993; Wallrabenstein et al., 1995; for review see Schink, 1997). Growth with propionate and sulfate was extremely slow. If simultaneously an H2-scavenging Desulfovibrio strain was present in sulfate-containing medium, the propionateoxidizing strains grew syntrophically by interspe-

719

cies H2 transfer rather than by utilizing sulfate themselves. Apparently, the pathway of sulfate reduction is poorly developed in the propionate oxidizers. Carbon Assimilation Heterotrophic Growth The organic compounds utilized as electron acceptors by sulfatereducing bacteria serve simultaneously as carbon sources for cell synthesis. Carbon dioxide is an important additional carbon source for various carboxylation reactions during biosynthesis. With several H2-utilizing species, the capacity for autotrophic growth with CO2 as the only carbon source was demonstrated (see next section). Hydrogen-utilizing sulfate-reducing bacteria of the genus Desulfovibrio, which are complete oxidizers, require acetate in addition to CO2 for cell synthesis (Mechalas and Rittenberg, 1960; Postgate, 1960; Sorokin, 1966a; Sorokin, 1966b; Sorokin, 1966c; Badziong et al., 1979; Brandis and Thauer, 1981; Brysch et al., 1987). Also species of the genera Desulfobulbus and Thermodesulfobacterium, and Desulfomicrobium norvegicum (formerly Desulfovibrio desulfuricans strain Norway 4) required acetate as organic carbon source (Brysch et al., 1987; F. Widdel, unpublished observation). The observation that approximately one third of cell carbon is derived from CO2 and two thirds derived from acetate (or molar CO2 : acetate = 1 : 1; Sorokin, 1966a; Sorokin, 1966b; Sorokin, 1966c) is explained by the pyruvate synthase reaction. Pyruvate synthase or pyruvate:ferredoxin oxidoreductase (PFOR), that carboxylates acetyl CoA reductively (acetyl-CoA + CO2 + 2 e- + 2H+ Æ 2 pyruvate + CoA) is a central metabolic enzyme (Badziong et al., 1979; Brandis-Heep et al., 1983; Schauder et al., 1987). The biosynthetic reaction is always required if the carbon source is acetate or a substrate that yields exclusively acetyl-CoA, e.g., ethanol or C-even fatty acids (or CO2 in autotrophs; see below). Hence, sulfate-reducing bacteria growing on ethanol or C-even fatty acids without the capacity for complete oxidation are expected to strictly require external CO2 for growth. It is unknown whether sulfate-reducing bacteria employ the same PFOR for acetyl-CoA assimilation and for pyruvate oxidation, e.g., during growth on lactate, or whether there are specifically regulated isoenzymes. Several further assimilatory enzymes have been studied in Desulfobacter species that employ a citric acid cycle for acetyl-CoA oxidation (Brandis-Heep et al., 1983; Schauder et al., 1987). In addition to pyruvate synthase, anaplerotic reactions include acetate activation via acetyl-CoA synthetase (acetate + ATP Æ acetyl-

720

R. Rabus, T.A. Hansen and F. Widdel

CoA + PPi; in addition to succinyl-CoA: acetate transferase), phosphoenolpyruvate (PEP) synthetase (pyruvate + ATP Æ PEP + AMP + Pi), and PEP carboxylase (PEP + HCO3- Æ oxaloacetate + Pi) to compensate for the withdrawal of a-ketoacids for biosynthesis. Phosphoenolpyruvate is also expected to serve for synthesis of triose and higher sugar phosphates. In incompletely oxidizing sulfate-reducing bacteria, similar reactions may provide PEP and oxaloacetate; further biosynthetic precursors can then be synthesized via sequences of an incomplete citric acid cycle. A citrate synthase with (R)-specificity has been studied in incompletely oxidizing Desulfovibrio species (Gottschalk, 1968). Autotrophic Growth When sulfate-reducing bacteria were shown to use H2 as an inorganic electron donor, also the possibility that cell synthesis might occur autotrophically from CO2 became of interest (Stephenson and Strickland, 1931). Carbon autotrophy was reported for sulfate-reducing enrichment cultures (Wight and Starkey, 1945) and Desulfovibrio strains (Butlin and Adams, 1947; Sisler and ZoBell, 1951). Later however, growth experiments and labeling studies with Desulfovibrio species revealed repeatedly that these sulfate reducers were lithoheterotrophs that required acetate in addition to CO2 for cell synthesis (Mechalas and Rittenberg, 1960; Postgate, 1960; Sorokin, 1966a; Sorokin, 1966b; Sorokin, 1966c; Badziong et al., 1979; Brandis and Thauer, 1981; Brysch et al., 1987). However, several newly isolated, completely oxidizing sulfate-reducing bacteria grew with H2 plus CO2 (or formate) in the absence of other carbon compounds (Widdel, 1980; Widdel and Pfennig, 1981; Pfennig et al., 1981c). Labeling studies with formate-utilizing Desulfoarculus (formerly Desulfovibrio) barsii (Jansen et al., 1984; 1985) and H2-utilizing Desulfobacterium species, Desulfobacter hydrogenophilus, Desulfosarcina variabilis, Desulfonema limicola and Desulfotosporosinus orientis (Brysch et al., 1987) clearly demonstrated the capacity for autotrophic growth. Several autotrophic strains excreted traces of acetate if incubated with H2 plus CO2 (or formate), with limiting sulfate concentrations. Enrichment cultures with H2 plus CO2 in sulfate-containing media were shown to yield mixed cultures of non-autotrophic Desulfovibrio species and autotrophic homoacetogenic bacteria, the latter providing acetate to the former (Brysch et al., 1987). Such mixed cultures grew faster than truly autotrophic sulfate-reducing bacteria; hence direct enrichment of the latter (autotrophic bacteria) from natural samples under autotrophic conditions is unlikely.

CHAPTER 1.22

With the exception of Desulfotosporosinus orientis, the facultative lithoautotrophic sulfate reducers are complete oxidizers (Brysch et al., 1987). Indeed, the mechanisms of CO2 fixation were found to be reverse reactions from the pathways which during organotrophic growth serve for acetyl-CoA oxidation. Whereas Desulfobacter hydrogenophilus assimilated CO2 via a reductive citric acid cycle (Schauder et al., 1987), Desulfobacterium autotrophicum used the inverse C1-pathway or Wood pathway (Schauder et al., 1989), viz. a sequence of reactions observed in homoacetogenic bacteria. The former has to reactivate acetate liberated in the succinyl-CoA:acetate CoA-transferase reaction. Formed acetyl-CoA from both pathways is then converted to pyruvate (see preceding section). Lithoautotrophically and organotrophically grown cells of D. autotrophicum exhibited different patterns of CO dehydrogenase aggregates during gel electrophoresis (Schauder et al., 1989). Obviously, the reductive and oxidative pathway, respectively, employed somewhat different enzymes. This indicates that formation of enzymes for the reductive and the oxidative pathways is regulated depending on whether H2 or organic electron donors are present. Desulfotosporosinus orientis can grow autotrophically but cannot oxidize organic substrates completely to CO2 (Klemps et al., 1985). The CO-dehydrogenase activity (R. Klemps and F. Widdel, unpublished observation) and a weak capacity for homoacetogenic growth (Klemps et al., 1985) suggests that this sulfate reducer also uses the C1-pathway for CO2 fixation. It is not understood why the assimilatory pathway in this species cannot be reversed for acetylCoA oxidation. Another incomplete oxidizer, Desulfomicrobium apsheronum also has been reported to grow autotrophically (Rozanova et al., 1988a). Assimilation of Nitrogen Compounds Ammonium represents the most readily used nitrogen source for sulfate-reducing bacteria and for other bacteria. Ammonium ions are common in anoxic habitats as a result of biomass degradation. In cultivation media for sulfate-reducing bacteria, ammonium salts are usually included. In sulfate reducers that can use nitrate as electron acceptor, its dissimilatory reduction to ammonium provides simultaneously a nitrogen source. Diazotrophic growth has been demonstrated in species of the genera Desulfovibrio (RiedererHenderson and Wilson, 1970; Lespinat et al., 1987; Postgate and Kent, 1985; Moura et al., 1987), Desulfobacter (Widdel, 1987), Desulfobulbus (Bomar M. and F. Widdel, unpublished

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

721

observation) and Desulfotomaculum (Postgate, 1970). The DNA carrying nifH/nifD hybridized with DNA from 13 diazotrophic strains of Desulfovibrio belonging to 5 different species; from D. gigas, the nifH gene coding for the Fe protein of the nitrogenase system was sequenced (Postgate et al., 1988; Kent et al., 1989).

metabolizing and fermentative extreme thermophiles of the Crenarchaeota. An overview of physiological properties of sulfate-reducing Archaeoglobus species is given in Table 7. Another member of this lineage is the hyperthermophilic archaeum Ferroglobus placidus, which can use thiosulfate as electron acceptor for the oxidation of H2 (Hafenbradl et al., 1996).

Sulfate-Reducing Archaea Archaeoglobus fulgidus was isolated from a submarine hydrothermal area and was identified as the first representative of the archaeal domain of life that could conserve energy via dissimilatory sulfate reduction (Stetter et al., 1987; Stetter, 1988; Zellner et al., 1989b). Two other Archaeoglobus species, A. profundus (Burggraf et al., 1990) and A. lithotrophicus (Stetter et al., 1993), are further archaeal sulfate reducers. A fourth Archaeoglobus species, A. veneficus, uses sulfite but not sulfate as electron acceptor (Huber et al., 1997). Archaeoglobus species typically grow optimally at temperatures above 80∞C and require at least 10 g NaCl/liter for growth (Stetter, 1992). Phylogenetic analyses revealed that the genus Archaeglobus represents a lineage within the Euryarchaeota (Woese et al., 1991) with particular relationships to methanogenic archaea; Archaeglobus is unrelated to the sulfur-

Reduction of Sulfate to Sulfide Transport of sulfate has not been studied so far in Archaeoglobus. The general pathway of sulfate reduction to sulfide in Archaeoglobus is analogous to the one established for sulfate-reducing bacteria (Dahl and Trüper, 1999b). The presence of the enzymatic activities essential for dissimilatory reduction of sulfate (ATP sulfurylase, APS reductase and sulfite reductase) were demonstrated in A. fulgidus (Speich and Trüper, 1988; Dahl et al., 1994). In Table 3, the sulfite-reductase from A. fulgidus is compares the sulfite-reductase from other prokaryotes mentioned. Activation of Sulfate Prior to reduction, sulfate is activated in an ATP-dependent reaction to APS, a reaction catalyzed by ATP sulfurylase. The dissimilatory ATP sulfurylase was purified from A. fulgidus and found to have a molecular weight of about 150 kDa (Dahl et al., 1988;

Table 7. Physiological properties of sulfate-reducing Archaeoglobus species.a Organic substrates utilized with SO42- and/or S2O32Lactate

Pyruvate

with S2O32-

+

nr

Formamide, glucose, starch, peptone, methanol, ethanol

-

+

nr

+

+

2,3-Butandiol, fumarate

-

+

nr

-

+

+

Valerate

-

-

+

nr

+c

+c

+c

-

-

nr

nr

nr

nr

+

nr

+ (obligate lithoheterotrophic) nr

Acetate

with SO42-

Formate

Others

Lithoheterotrophic (+acetate) with SO42-

Temp. Opt. [∞C] 83

+

nr

75–80

+

-

A fulgidus strain 7342b

76

-

A profundus strain AV18b

82

A lithotrophicus

80

Species A fulgidus strain VC-16b

A fulgidus strain Zb

H2 utilization Lithoautotrophic

Symbols: +, utilized; -, not utilized; nr, not reduced. a Species of the genus Archaeglobus are the only sulfate-reducing Archaea known so far. A veneficus strain SNP6 does not reduce sulfate, even though this species is capable of sulfite and thiosulfate reduction (Huber et al., 1997). b Data obtained from: A fulgidus strain VC-16 (Stetter, 1988), A fulgidus strain Z (Zellner et al., 1989b), A fulgidus strain 7342 (Beeder et al., 1994), A profundus strain AV18 (Burggraf et al., 1990), A lithotrophicus (Stetter et al., 1993). c Utilization strictly dependent on the presence of H2. It is presently unknown whether these compounds are co-metabolically utilized as electron donors, or only as carbon sources.

722

R. Rabus, T.A. Hansen and F. Widdel

1990). The coding gene for sulfate adenylyltransferase (sat) was cloned and found to exhibit homology with the coding genes of homooligomeric ATP sulfurylases from various bacteria and eukaryotes. The sat gene was cloned and overexpressed in E. coli and the recombinant protein was purified. It was found to be a homodimer. Activity testing proved that the recombinant protein could indeed form ATP from APS and PPi (Sperling et al., 1998; Sperling et al., 1999). Reduction of APS The enzyme APS reductase catalyzes the two-electron reduction of APS to sulfite and AMP. The enzyme was purified from A. fulgidus and characterized. An apparent molecular mass of 160 kDa was determined and the protein was found to contain one FAD and [FeS] clusters (Speich and Trüper, 1988; Dahl et al., 1994). Spectroscopic studies of the purified enzyme demonstrated that the enzyme contained two distinct [4Fe-4S] clusters which showed similarity to the ones identified in the APS reductase from Desulfovibrio gigas (Lampreia et al., 1991). Analysis of the purified APS reductase on SDS-PAGE revealed two bands corresponding to molecular masses of 80 kDa and 18.5 kDa. Taking the apparent molecular mass of the holoenzyme into account, this finding suggested a a2b structure for the enzyme. The presence of two different subunits was confirmed by the analysis of the genes coding for the a- and b-subunit, aprA and aprB, respectively. The aprA and aprB genes encoded a 73.3 kDa and a 17.1 kDa polypeptide, respectively. The aprA gene product showed homologies to flavoproteins from Escherichia coli and Bacillus subtilis, whereas the aprB gene contained sequences for cysteine clusters that could ligate the [FeS] centers identified by the spectroscopic analyses (Speich et al., 1994). Reduction of Sulfite The six-electron reduction of sulfite to sulfide is catalyzed by the sulfite reductase. This enzyme was purified from A. fulgidus and exhibited characteristics similar to those of dissimilatory sulfite reductases from other bacteria. The native enzyme had an apparent molecular mass of 218 kDa and consisted of two subunits with molecular masses of 40 and 50 kDa, suggesting a a2b2 structure. Furthermore, the holoenzyme contained two sirohemes and six [4Fe-4S] clusters. The genes encoding the a- and b-subunit, dsrA and dsrB, were cloned, and found to be arranged in an operon structure. The deduced DsrA peptide contains the cysteine residues required for the coordination of siroheme-[4Fe-4S] complexes. Furthermore, both deduced peptides, DsrA and DsrB, contain addi-

CHAPTER 1.22

tional cysteine residues which are characteristic of binding motifs for ferredoxin-like [4Fe-4S]clusters. Thus the findings of the sequence analyses corroborated the biochemical data directly obtained from the purified protein. The dsrA and dsrB genes showed a high degree of similarity suggesting that these genes arose by duplication from an ancestral gene. Comparative sequence analyses of sulfite reductases from various microorganisms revealed that only sulfite reductases from A. fulgidus and Salmonella typhimurium contained a ferredoxin-like domain in the proximity of the conserved putative siroheme-[4Fe4S]-binding cysteine residues (Dahl et al., 1993; 1994). The dsrAB genes of A. fulgidus were also highly homologous to the dsvAB genes that code for desulfoviridin of Desulfovibrio vulgaris (Karkhoff-Schweizer et al., 1995). A dissimilatory sulfite reductase was also isolated from the hyperthermophilic archaeon Pyrobaculum islandicum. This archaeon cannot reduce sulfate; however, it is capable of organotrophic growth with sulfite as electron acceptor (Huber et al., 1987). The purified sulfite reductase was shown to have a a2b2 structure and to contain siroheme and [FeS] clusters. Two coding genes (dsrA and dsrB) could be cloned and found to be organized in an operon. Downstream of the dsrB gene, a third gene, dsrC, was identified which was homologous to the proposed g-subunit of the sulfite reductase from Desulfovibrio vulgaris (Molitor et al., 1998; Dahl et al., 1999a). Electron Acceptors Other Than Sulfate In addition to sulfate, A. fulgidus can utilize thiosulfate and sulfite as electron acceptor. The utilization of sulfite is understandable because it is an intermediate during sulfate reduction. The reduction of thiosulfate in A. fulgidus has not been studied in more detail. Electron Carriers Ferredoxin Ferredoxin is an electron carrier which has been frequently encountered in sulfate-reducing bacteria. It has also been identified in A. fulgidus. Ferredoxin is involved in catabolic reactions in A. fulgidus, such as pyruvate:ferredoxin oxidoreductase (Kunow et al., 1995) and the acetyl-CoA decarbonylase/synthase (COdehydrogenase-containing) complex (Dai et al., 1998), and possibly also in pyruvate synthesis from acetyl-CoA in lithoheterotrophic species that use acetate as organic carbon sources. Menaquinone Tindall et al. (1989) discovered a novel menaquinone in A. fulgidus. This menaquinone possesses a fully saturated hepta-

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

prenyl side chain (MK-7H14) and is the major lipoquinone in A. fulgidus. Metabolism of Electron Donors Archaeoglobus species may grow chemolithoautotrophically with H2 and CO2, chemoorganotrophically on formamide, lactate, pyruvate, glucose and complex organic substrates (starch, peptone), or lithoheterotrophically on H2 and acetate, lactate, pyruvate or other organic compounds (Stetter, 1992). An overview of the metabolism of electron donors by Archaeoglobus species is given in Table 7. The occurrence of Archaeoglobus species in marine and terrestrial oil-field waters has been reported several times (Stetter et al., 1993; Beeder et al., 1994; L’Haridon et al., 1995) and has suggested that Archaeoglobus species may utilize constituents of crude oil. However a utilization of hydrocarbons, the main constituents of crude oil, could not be demonstrated. Lactate, Pyruvate, and Acetate A. fulgidus completely oxidizes lactate to CO2 with sulfate as electron acceptor (Möller-Zinkhan et al., 1989; Zellner et al., 1989b). Lactate is oxidized to acetyl-CoA via lactate-dehydrogenase and pyruvate:ferredoxin oxidoreductase (PFOR; Möller-Zinkhan et al., 1989). Based on its predicted function as lactate dehydrogenase, a gene (dld) was cloned from the completely sequenced genome of A. fulgidus (Klenk et al., 1997) and heterologously overexpressed in Escherichia coli. The purified recombinant protein possessed D-lactate dehydrogenase activity, contained Zn2+ and the flavin cofactor FAD (Reed and Hartzell, 1999). The PFOR has been purified from A. fulgidus and found to have an apparent molecular mass of 120 kDa, a heterotetrameric (abgd) structure and to contain thiamine pyrophosphate and iron-sulfur clusters (Kunow et al., 1995). Further oxidation of acetyl-CoA to CO2 proceeds via a C1/CO-dehydrogenase pathway that may be regarded as an archaeal analogue of the pathway in sulfate-reducing bacteria (Fig. 11B). A unique characteristic of the archaeal pathway is the involvement of the cofactors F420, tetrahydromethanopterin and methanofuran that had been detected before in methanogenic archaea (Stetter et al., 1987; Möller-Zinkhan et al., 1989; Möller-Zinkhan and Thauer, 1990). The CO dehydrogenase is part of a multienzyme complex termed acetyl-CoA decarbonylase synthase (ACDS) that was isolated and characterized (Dai et al., 1998). This multienzyme complex consists of five different subunits ranging from 18.5 to 89 kDa in molecular mass and catalyzes the cleavage of acetyl-CoA into a bound methylgroup and bound CO, or the reverse reaction. The methyl carrier is tetrahydromethanopterin

723

(H4MPT); also, tetrahydrosarcinopterin reacts as methyl carrier with the complex. Ferredoxin is employed as electron carrier by this multienzyme complex. Prior to the study presented by Dai et al. (1998), ACDS complexes had been detected only in methanogens. Structural and functional properties of the ACDS complex from A. fulgidus are similar to those of the complex from methanogens. Therefore much insight into the function of the ACDS complex in A. fulgidus is based on the studies of this complex in the methanogens. The complex from Methanosarcina barkeri, which has been studied best, also consists of five subunits and has a (abgd)6 structure, giving rise to the remarkable total molecular mass of ca. 2.0 MDa for the entire complex (Grahame, 1991). Carbon monoxide and CO2 can be used for carbonylation of methylated tetrahydrosarcinopterin. A hydrogenase that was resolved from the multienzyme complex was capable of reconstituting the acetyl-CoA synthesis of the complex (Grahame and DeMoll, 1995). Separation of the ACDS complex from Methanosarcina by limited proteolytic digestion allowed specific catalytic functions to individual subunits: the CO-dehydrogenase reaction is performed by the a component; the methyltransferase is located on the g-subunit and parts of the dsubunit; and the binding of CoA or acetyl-CoA occurs on the b-subunit (Grahame and DeMoll, 1996). The CH3-group from acetyl-CoA cleavage in Archaeoglobus fulgidus is further oxidized to CO2 via N5,N10-methylene-H4MPT reductase, N5,N10-methylene-H4MPT dehydrogenase, N5,N10-methenyl-H4MPT cyclohydrolase, formylmethanofuran:H4MPT formyltransferase and formylmethanofuran dehydrogenase (MöllerZinkhan et al., 1989). Purification of the corresponding enzymes from A. fulgidus allowed the C1-pathway of methyl oxidation to be unequivocally established and demonstrated that the enzymes from A. fulgidus had very similar molecular and catalytic properties as those of the acetate-degrading methanogens (Schmitz et al., 1991; Klein et al., 1993; Schwörer et al., 1993). Factor F420 serves as H2 acceptor for N5,N10methylene-H4MPT reductase and N5,N10methylene-H4MPT dehydrogenase. The natural electron acceptor for formylmethanofuran dehydrogenase is unknown. A membrane-bound F420H2:quinone oxidoreductase complex was purified from A. fulgidus. This enzyme complex is presumed to be involved in the chemiosmotic conservation (Kunow et al., 1994). Similarities in the enzymes and cofactors of the C1-pathway in Archaeoglobus and methanogens suggest a metabolic relationship. Indeed, Archaeoglobus was suggested to represent a link between hyperthermophilic sulfur-reducing (nonsulfate-reducing) and methanogenic

724

R. Rabus, T.A. Hansen and F. Widdel

archaea. However, Archaeoglobus does not possess the cofactors (mercaptoethanesulfonate, mercaptoheptanoyl threonine phosphate) and enzymes (methyltransferase, methyl-CoM reductase, heterodisulfide reductase) that are involved in the terminal step of CH4 formation from the H4MPT-bound methyl group. The formation of low amounts of CH4 observed in Archaeoglobus (Stetter et al., 1987) (and in sulfate-reducing bacteria; Schauder et al., 1986) is a by-reaction of the methyl group transferred by CO dehydrogenase. Malate, Isocitrate and Glutamate Even though Arc-

haeoglobus performs oxidation of acetate via the C1-pathway and not via the TCA cycle, activities of malate dehydrogenase and isocitrate dehydrogenase were measured in cell extracts of (Möller-Zinkhan et al., 1989). These enzymes presumably function in biosynthesis (Langelandsvik et al., 1997; Steen et al., 1997). Both enzymes were purified and found to possess pronounced thermostability. Malatedehydrogenase was specific for NAD+, whereas isocitrate dehydrogenase has a high preference for NADP+. Also a thermostable NADP+-specific glutamate dehydrogenase was purified from this archaeon. This enzyme accounts for 0.8% of the total cell extract protein, which is relatively large in view of the assumed function in the assimilation of ammonia (Aalén et al., 1997). Autotrophic Growth Autotrophic growth on H2 and sulfate as energy source and CO2 as carbon source was studied in Archaeoglobus lithotrophicus. All enzymatic activities and coenzymes required for the fixation of CO2 via the reductive CO dehydrogenase pathway were demonstrated in cell extracts of A. lithotrophicus (Vorholt et al., 1995). This reductive CO dehydrogenase pathway is, in principle, the reverse of the oxidative C1/CO-dehydrogenase pathway employed by A. fulgidus for the oxidation of acetyl-CoA. The same study by Vorholt et al. (1995) showed that CO dehydrogenase was lacking in A. profundus, which explained why this archaeon requires acetate for biosynthesis during growth on H2. Genome A. fulgidus strain VC-16 is the first sulfate- and sulfur-reducing microorganism and the second archaeon after Methanococcus jannaschii (Bult et al., 1996) the complete genomic sequence of which has been determined (Klenk et al., 1997). The genome consists of a single chromosome of about 2.2 Mb and has an average G+C content of 48.5 mol%. A total of 2,436 ORFs were iden-

CHAPTER 1.22

tified with an average size of 822 bp. Putative functions could be assigned to 1,797 ORFs, whereas the remaining 639 ORFs had no database matches. Two thirds of these unidentified ORFs are shared with M. jannaschii. Of the ORFs with assigned function, 719 genes can be classified into 242 families. The largest of these families is the superfamily of ATP-binding subunits of ABC transporters, which comprises 40 members in A. fulgidus. The genome of A. fulgidus contains three regions of short repeats (>40 bp), which are similar to those found in M. jannaschii, and nine classes of long repeated sequences ( 6 was shown to exceed 10 mM (Schauder and Müller, 1993), which is close to the apparent KM (about 20 mM) determined for polysulfide respiration in W. succinogenes (Klimmek et al., 1998). Thus there is evidence for the use of polysulfide as the actual electron acceptor in sulfur respiration. In the study of sulfur reduction, it must not be forgotten that other bacteria grow with elemental sulfur without the possibility of solubilization in the form of polysulfide. These are anaerobes that disproportionate elemental sulfur in the presence of sulfide-scavenging ferric minerals (Thamdrup et al., 1993) or aerobic bacteria that oxidize extracellular sulfur. It is unknown how these bacteria cope with the low solubility of sulfur in water. In photoautotrophic purple bacteria and possibly in aerobic sulfide oxidizers forming intracellular sulfur globules as intermediates, the sulfur is topologically periplasmatic (“extracytoplasmatic”) and associated with proteins; these complexes are assumed to control formation or further oxidation of the sulfur (Dahl, 1999b). Examples of other microorganisms growing with insoluble substrates are bacteria reducing ferric minerals, or bacteria or yeasts oxidizing long-chain alkanes. Ferric minerals

are probably reduced in direct contact with the cells (Lovley, 1995a), or by an extracellular cytochrome (Seeliger et al., 1998). Long-chain alkanes are also utilized in direct contact with the cells, or via pseudosolubilization with biotensides (Bühler and Schindler, 1984). Research on Wolinella succinogenes Wolinella was originally isolated as a fumarate-reducing bacterium utilizing H2 or formate as electron acceptor. The capacity for microaerobic growth has been formerly mentioned but not the subject of more recent studies. Wolinella was then shown to reduce sulfur (Macy et al., 1986), like Sulfurospirillum deleyanium, the former spirillum 5175 (Wolfe and Pfennig, 1977), a facultative microaerophile. In addition to H2 or formate, both sulfur reducers oxidize some other organic compounds such as lactate. Oxidation is incomplete and leads to acetate. Wolinella and Sulfurospirillum are not only metabolically, but also phylogenetically related; they belong to the e Proteobacteria. The Sud Protein In a study in which the involvement of polysulfide in sulfur respiration of Wolinella succinogenes was questioned, Fe2+ [as Fe(OH)2] was added to the medium to precipitate all sulfide formed by W. succinogenes as FeS and thus prevent formation of polysulfide. Under these conditions, W. succinogenes still grew anaerobically with formate and elemental sulfur, indicating that sulfur reduction is in principle possible without the involvement of polysulfide as an intermediate (Ringel et al., 1996). From the iron(II)-containing culture of W. succinogenes, a soluble sulfur-containing fraction was isolated that by treatment with CN- could be separated further into a yet unidentified sulfur species and the so-called Sud protein (Hedderich et al., 1999). The coding sud gene was isolated from W. succinogenes and its sequence indicated the presence of a signal peptide and only one cysteine in the polypeptide chain. The recombinant Sud protein was purified after heterologous expression in Escherichia coli. The enzyme consists of two identical subunits (14 kDa), lacks any prosthetic groups or heavy metals and is located in the periplasm. The synthesis of the Sud protein is induced during growth on elemental sulfur and polysulfide (Kreis-Kleinschmidt et al., 1995). Further studies with a His-tagged Sud protein (Sud-His6), which was also purified from E. coli, demonstrated a catalysis of the formation of thiocyanate from cyanide and polysulfide with an apparent KM of less than 20 mM polysulfide. The monomer of Sud-His6 was found to bind up to 10 sulfur atoms from polysulfide. Addition of small amounts of Sud-His6 to membrane fractions of

CHAPTER 1.22 Fig. 18. Possible generation of a proton-motive force (pmf) during growth of Wolinella succinogenes or other spirilloid sulfur reducers on H2 and sulfur. Diffusion and collision of HydC and PsrC is assumed to be required for electron transfer. The mechanism for generation of a proton gradient is not known. Possibly, protons are translocated via proteinbound menaquinone to the periplasm. Abbreviations: HydABC, subunits of hydrogenase; PsrABC, subunits of polysulfide reductase; Sud, protein that increases the availability of polysulfide (formerly termed “sulfide dehydrogenase”). [S] indicates a soluble form of sulfur, most probably polysulfide. The scheme was adapted from Hedderich et al. (1999).

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes + + +

Periplasm

OM

H2

HydB Ni

2

HydA 2 e– Fe/S

H+

CM

– – –

727

Cytoplasm

HydC Cytb 2 e–

H+ + [S] Sud HS–

H+ Mo PsrA

2 e– Fe/S

MKH– MK

PsrB

PsrC

H+

1 / ATP 3

+ + +

W. succinogenes stimulated the electron transport from H2 to polysulfide (Klimmek et al., 1998). A deletion mutant of the sud gene (Dsud) was constructed in W. succinogenes by homologous recombination. However, growth of the Dsud deletion mutant on formate and polysulfide as compared to that of the wild type was not affected (Kotzian et al., 1996). By site directed mutagenesis, the single cysteine residue in the Sud protein (Cys109) was replaced by a serine residue. The modified Sud protein (C109S)SudHis6 showed marked differences from the Sud-His6 protein. The (C109S)Sud-His6 protein neither catalyzed formation of thiocyanate from cyanide and polysulfide nor stimulated the electron transport to polysulfide. Moreover, the Cys109 residue was found to be required for binding polysulfide-sulfur to the Sud protein. Despite some inconsistent results from growth experiments, the Sud protein is assumed to function in transferring sulfur from aqueous polysulfide to the active site of polysulfide reductase (Klimmek et al., 1999). The Sud protein and polysulfide reductase (Psr) were present in nearly equimolar amounts when W. succinogenes was grown on polysulfide, and it is assumed that Sud is bound to Psr (Fig. 18; Hedderich et al., 1999). Polysulfide Reductase The enzyme that catalyzes the reduction of polysulfide sulfur to sulfide is termed “polysulfide reductase” (Psr). This

– – –

1 / ADP 3 + Pi

enzyme is encoded by the polysulfide reductase operon (psrABC; Krafft et al., 1992). The nucleotide sequence of the psrABC genes indicates that Psr is a heterotrimer consisting of three subunits (PsrA, B and C). The PsrA (81 kDa) and PsrB (21 kDa) subunits are hydrophilic proteins, whereas PsrC (34 kDa) is of hydrophobic nature with eight putative transmembrane-spanning segments and is assumed to function as membrane anchor of the Psr holoenzyme. The PsrA subunit was found to be homologous to known molybdoenum-containing oxidoreductases, such as formate dehydrogenase of E. coli. Indeed, a molybdopterin guanine dinucleotide was identified in the purified protein (Jankielewicz et al., 1994). The PsrA subunit is assumed to be the catalytic subunit of the Psr protein. The psrA gene also includes the coding sequence for a leader peptide indicating an orientation of the PsrA subunit toward the periplasm. Moreover, PsrA was identified in the periplasmic fraction of a DpsrC mutant. Based on the predicted presence of 16 cysteine residues, PsrB is assumed to contain several [FeS] clusters involved in electron transfer. Even though the psrB gene does not contain a coding sequence for a leader peptide, a periplasmic orientation of PsrB is postulated (Krafft et al., 1992). The purified Psr contains 1 mol menaquinone per mol of protein. Menaquinone is assumed to serve as acceptor for electrons transferred from hydrogenase and as direct electron donor for polysulfide/sulfur reduction.

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The purified Psr protein catalyzes the reduction of polysulfide to sulfide with BH4- as hydride donor, and the oxidation of sulfide to polysulfide by 2,3-dimethyl-1,4-naphthoquinone, a soluble analogue of menaquinone (Krafft et al., 1992). In wild type W. succinogenes, polysulfide reductase activity is still present and active when cells are grown with fumarate as electron acceptor (Lorenzen et al., 1993). Deletion mutants of Psr (DpsrC, DpsrBC and DpsrABC) were grown with fumarate and cell fractions were analyzed for their capacity to oxidize sulfur and to transfer electrons from formate to polysulfide (Krafft et al., 1995). The DpsrC mutant catalyzed the oxidation of sulfide with dimethylnaphthoquinone, which was not observed with the DpsrABC or DpsrBC mutant. This indicated that PsrA and PsrB, but not PsrC, were directly involved in the transfer of reducing equivalents to a quinone site. However, the capacity of the DpsrC mutant to perform the entire electron transfer from formate to polysulfide was only 5% of the wild type activity, suggesting that PsrC is required for further electron transport reactions. If the DpsrABC mutant was grown on polysulfide instead of fumarate, activity for sulfide oxidation and polysulfide reduction could still be measured. A so far unidentified protein could be extracted from the membranes of the polysulfide-grown mutant that seems to replace polysulfide reductase. Electron Transport from Formate or H2 to Polysulfide Wolinella succinogenes utilizes either H2 or formate as electron donors (Macy et al., 1986). The same hydrogenase and formate dehydrogenase are operative if either sulfur or fumarate are used as electron acceptors (Schröder et al., 1988). A hydrogenase deletion mutant (DhydABC) did not grow with H2 and polysulfide, or with H2 and fumarate. Growth could be restored by complementing the mutant with the hydABC operon (Gross et al., 1998a; Gross et al., 1998b). Electrons from hydrogenase and formate-dehydrogenase have to be transferred to polysulfide reductase. Electron transfer reactions were most intensely studied with formate. Even though the substrate-binding sites of formate dehydrogenase and hydrogenase are both orientated toward the periplasm (Kröger and Winkler, 1981), formate does not diffuse through membrane bilayers and thus allows more defined studies in vesicles than H2. Electron transfer from formate was studied in vesicles as a function of the ratio between phospholipid and membrane proteins, by dilution of the membrane fraction of W. succinogenes with phospholipid. Based on these experiments, a model of

CHAPTER 1.22

diffusion and collision was suggested. Collision of hydrogenase or formate dehydrogenase, respectively, with the polysulfide reductase is regarded as a requirement for this electron transfer. In addition to the collision of proteins, menaquinone bound to PsrC is essential (Hedderich et al., 1999). In contrast, electron transport to fumarate reductase in the cytoplasmic membrane of W. succinogenes does not involve direct collision of proteins but rather occurs via freely diffusible menaquinone in the cytoplasmic membrane (Jankielewicz et al., 1995; Hedderich et al., 1999). Properties of hydrogenase in W. succinogenes have been studied in detail. The enzyme is membrane-bound, contains nickel, and catalyzes the reduction of dimethylnaphthoquinone or benzylviologen with H2 (Unden et al., 1982). It could be isolated from the membrane fraction of W. succinogenes and was found to consist of three subunits, HydA (30 kDa), HydB (60 kDa) and HydC (23 kDa). A deletion mutant without the hydrogenase (DhydABC) cannot grow with H2 and either polysulfide or fumarate. The three subunits of hydrogenase are encoded by three adjacent genes, hydABC. The HydA subunit is a hydrophilic protein that is likely to be localized in the periplasm because the gene, hydA, contains a coding sequence for a leader peptide. The HydA subunit contains eight cysteine residues, some of which are possible ligands for [FeS] clusters. The C-terminus of HydA contains about 20 hydrophobic residues that could constitute a membrane anchor by forming a transmembrane helix and in this way a membrane anchor for the protein. The HydB protein, the catalytic subunit of hydrogenase, is hydrophilic and contains eight cysteine residues that are likely to coordinate [FeS] clusters. The Cys546 residue is possibly functioning in ligation of Ni. The catalytic subunit HydB of the intact hydrogenase is located in the periplasm as demonstrated with activity tests and western blot analyses of cell fractions. The HydA and HydB proteins are homologous to the corresponding subunits of other known Ni-hydrogenases. The HydC subunit is a hydrophobic protein with four putative transmembrane-spanning segments. Biochemical studies indicated that HydC represents a cytochrome b, with the two heme-B groups ligated by four His residues. Mutants created by substitution of the heme-ligating His residues no longer had the activity to reduce quinone with H2 and to transfer electrons to polysulfide reductase. These results indicate that the menaquinone bound as a prosthetic group to the PsrC is the primary acceptor for electrons from cytochrome b of HydC. This finding supports the assumption that also hydrogenase has

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

to be associated with polysulfide reductase for electron transfer in the membrane (Dross et al., 1992; Gross et al., 1998b) as in the case of formate dehydrogenase. The exact mechanism for the generation of the electrochemical proton gradient with formate or H2 as electron donors is not known. Possibly PsrC couples electron transfer via bound menaquinone to polysulfide to a translocation of protons (Hedderich et al., 1999; Fig. 18). Polysulfide and fumarate respiration in W. succinogenes differ not only with respect to the involvement of quinone. Also, the orientation of the two reductases is different. Whereas the substrate-binding site of polysulfide reductase is oriented toward the periplasm, that of fumarate reductase is localized on the cytoplasmic side of the membrane (Kröger et al., 1980). The substrate-binding sites of hydrogenase and of formate dehydrogenase both face the periplasm (Kröger and Winkler, 1981). Regulation of Sulfur Respiration Growth cultures of W. succinogenes on sulfur and formate in medium that also contained nitrate or fumarate, reduced sulfur but neither of the other two electron acceptors. This indicated that the energetically less favorable electron acceptor, sulfur, represses the utilization of the more favorable electron acceptors. In contrast, cells that were grown with nitrate or fumarate could respire both of these electron acceptors. Polysulfide reductase activity in fumarate-grown cells was as high as in sulfur-grown cells, but rather low in nitrate-grown cells (Lorenzen et al., 1993). In conclusion, regulation of anaerobic respiration with alternative electron acceptors is not clearly in accordance with their energetic “hierarchy.” Electron Acceptors Other Than Sulfur Wolinella succinogenes also can grow with nitrate and fumarate as electron acceptors. Nitrate is reduced to ammonia and not to N2 as in “true” denitrifying bacteria. A hexaheme cytochrome c3 acting as nitrite reductase has been isolated from Wolinella succinogenes (Liu et al., 1983). Another nitrogen compound reduced by Wolinella succinogenes is N2O; unlike nitrate (or nitrite), N2O is reduced to N2 (Yoshinari, 1980). Furthermore, spirilloid sulfur reducers closely related to Wolinella succinogenes were shown to reduce dimethylsulfoxide to dimethylsulfide (Zinder and Brock, 1978; Widdel, 1988). In connection with the initial characterization of Wolinella succinogenes, microaerobic growth has been reported (Wolin et al., 1961). Microaerobic growth was also shown in related spirilloid sulfur-reducing bacteria (Wolfe and Pfennig, 1977; Widdel, 1988).

729

Research on Desulfuromonas and Desulfurella Among the sulfur-reducing bacteria, the capacity for a complete oxidation of organic substrates occurs necessarily in those genera and species that grow on acetate (Desulfuromonas and Desulfurella). Desulfuromonas (Pfennig and Biebl, 1976) and Desulfurella (Bonch-Osmolovskaya et al., 1990) were directly isolated with acetate and sulfur. Both are obligate anaerobes. They are members of the d Proteobacteria, with a specific relationship of Desulfuromonas to completely oxidizing sulfate-reducing bacteria. In addition to acetate, Desulfuromonas species may utilize a number of simple organic compounds (Table 2). Oxidation of Acetate via the Citric Acid Cycle The presence of all enzymes of the citric acid cycle could be demonstrated in Desulfuromonas acetoxidans and Desulfurella acetivorans (Gebhardt et al., 1985; Schmitz et al., 1990; for overview see Kröger et al., 1988; Thauer, 1988; Thauer, 1989a; Thauer et al., 1989b). [14C]-labeling experiments demonstrated a functioning citric acid cycle in Desulfuromonas acetoxidans (Gebhardt et al., 1985). Desulfuromonas activates acetate like Desulfobacter via succinyl-CoA:acetate CoA transferase (Fig. 12). In Desulfurella acetivorans, however, the formation of succinate from succinyl-CoA is associated with the synthesis of one ATP; this amount is used again to activate acetate by acetate kinase (Schmitz et al., 1990; Fig. 12 C). Citrate formation in Desulfuromonas acetoxidans occurs with si-specificity, as in Desulfobacter, but without coupling to ATP formation. Malate in Desulfuromonas and Desulfurella is dehydrogenated with NAD+, as in mitochondria and most bacteria; Desulfurella has in addition NADP+specific malate dehydrogenase. The reduction of NAD(P)+ (E0¢ = –0.32 V) with malate (E0¢ = –0.166 V) in the sulfur reducers is understood in view of the way of citrate synthesis. By not being coupled to ATP formation, the reaction is exergonic and “pulls” the energetically unfavorable dehydrogenation of malate with pyridine nucleotides. Furthermore, an NADP:ferredoxin oxidoreductase has been detected (Kröger et al., 1988). A comparison of the modifications of the citric acid cycle found in sulfate- and sulfur-reducing bacteria is presented in Fig. 12. Inasmuch as neither Desulfuromonas nor Desulfurella gain net ATP by substrate-level phosphorylation, energy conservation must be achieved by chemiosmosis. The electron transport from ferredoxin, which might accept electrons from 2-oxoglutarate and via (NADP), from

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isocitrate, to the postulated electron donor for the sulfur reductase (DE’ around 0.2 V) could pump 2H+/2e- or 4H+/acetate. As in Desulfobacter, succinate oxidation in Desulfuromonas acetoxidans to fumarate (E0¢ = +0.033 V) with menaquinone (E0¢ = –0.074 V) is endergonic from the viewpoint of standard potentials. It appears unlikely that the reaction is made feasible solely by shifting concentration ratios of involved redox couples, or by specific coupling to a favorable redox reaction (see Desulfobacter), because electrons from menaquinone have to be transported further to sulfur reductase (S/H2S, E¢ = –0.19 V for 10 mM H2S; Fig. 14B). It is more likely that electron transport from succinate (oxidation) to sulfur (reduction) with a redox span of DE¢ = –0.22V is driven by chemiosmosis (reversed electron transport). With the consumption of 2 H+ for the energy-driven oxidation of 1 succinate, 2 H+/acetate remain for ATP synthesis, yielding 1/2 to 2/3 mol ATP/mol acetate. This is in relatively good agreement with energetic considerations based on growth yields. The growth yield of Desulfuromonas acetoxidans growing on acetate and sulfur was 4.2 g dry mass/mol acetate at an average doubling time of 3.8 hours (Pfennig and Biebl, 1976). The yield of Desulfovibrio vulgaris at this doubling time with acetate as C-source (H2 as electron donor) was 9.1 g/mol sulfate (Nethe-Jaenchen and Thauer, 1984). Assuming a similar maintenance and YATP, Desulfuromonas acetoxidans should have a net yield of around 0.6 mol ATP/mol acetate. Cytochromes Sulfur-reducing bacteria of the genus Desulfuromonas contain large amounts of various cytochromes (Pfennig and Biebl, 1976; Bache et al., 1983). A triheme c-type cytochrome, referred to as cytochrome c551.5 or c7, has been characterized (Probst et al., 1977; Fiechtner and Kassner, 1979). A c-type cytochrome has been suggested to transport electrons to sulfur reductase in Desulfuromonas (Kröger et al., 1988). Indeed, the cytochrome c551Ÿ.5 was demonstrated to reduce polysulfide in Desulfuromonas acetoxidans, which is indicative of its function in terminal reduction (Pereira et al., 1997). In addition, the cytochrome c551Ÿ.5 from Desulfuromonas acetoxidans was shown to function in Fe(III) reduction (Roden and Lovley, 1993; Lojou et al., 1998); the final reduction of insoluble Fe(III)minerals must occur in direct contact with the cell and therefore requires electron transport through the cytoplasm. The structural gene of cytochrome c551Ÿ.5 was cloned and heterologously overexpressed in Desulfovibrio desulfuricans. The purified recombinant cytochrome c551Ÿ.5 had the same biochemical and metal-reducing properties as the protein from Desulfuromonas acetoxidans (Aubert et al., 1998b). Structural

CHAPTER 1.22

analysis revealed strong analogies between the triheme cytochrome c551Ÿ.5 and the tetraheme cytochrome c3. The region that harbors the heme II group in c3 is not present in c551Ÿ.5. However, the orientation of the other three heme groups is very similar in the two cytochromes (Banci et al., 1996; Coutinho et al., 1996; Turner et al., 1997). Recently two new c-type cytochromes were isolated from Desulfuromonas acetoxidans, a monoheme cytochrome c (M = 10 kDa) and a tetraheme cytochrome c (M = 50 kDa), both of which are located in the periplasm (Bruschi et al., 1997). Hexaheme and octaheme cytochromes also have been isolated from Desulfuromonas acetoxidans (Pereira et al., 1997). Sulfur-Reducing Archaea The capacity to reduce elemental sulfur to sulfide is found in several genera of hyperthermophilic archaea (Stetter, 1996; Hedderich et al., 1999). If growth occurs with H2 as electron donor (+CO2), energy conservation can be only explained by a chemiosmotic process rather than by substrate-level phosphorylation. If such prokaryotes utilize alternatively organic compounds as electron donors (and carbon sources), one may assume that also the metabolism of these substrates involves chemiosmotic energy conservation during sulfur reduction. This section will primarily treat such sulfur-respiring or “true” sulfurreducing archaea, as far as their biochemistry has been investigated. In addition, results from investigations on Pyrococcus furiosus are included; fermentative growth of this archaeon is stimulated by sulfur. Biochemistry of this species has been investigated intensively. Reduction of Sulfur and Polysulfide Archaeons of the genus Pyrodictium grow chemolithotrophically by sulfur respiration at around 100∞C (Fischer et al., 1983; Stetter et al., 1993). A H2:sulfur-oxidoreductase complex was isolated from the membrane fraction of Pyrodictium abyssi isolate TAG11. This enzyme complex was shown to consist of nine polypeptides with an estimated total molecular mass of 520 kDa. The enzyme complex contains several uncharacterized [FeS] clusters, Ni and Cu ions, two cytochrome b and one cytochrome c. The enzyme complex is proposed to encompass hydrogenaseand sulfur-reductase activity as well as electron carrier components; the molecular arrangement is supposed to allow the coupling of S0 reduction with H2 to energy conservation (Dirmeier et al., 1998). The organization of the different components in such a large enzyme complex may allow stabilization of the interacting components and represent a strategy in hyperthermophiles to

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

perform sulfur respiration at temperatures of around 100∞C. Pyrococcus furiosus grows at 100∞C by fermentation of carbohydrates to acetate, CO2 and H2. If sulfur is present in the medium, H2S is produced in addition to H2, and the growth yield increases (Fiala and Stetter, 1986; Schicho et al., 1993). There are doubts whether sulfur reduction is a respiratory, chemiosmotically coupled process. Sulfur may serve as an electron sink for certain dehydrogenations and render fermentation more effective. Growth with H2 + sulfur as energy source has not been observed. An “H2evolving” hydrogenase was purified from P. furiosus which had a heterotrimeric (abg) structure and contained one [2Fe-2S] cluster and Ni (Bryant and Adams, 1989). Polysulfide can be reduced to H2S by this hyperthermophile and is assumed to be the natural substrate during sulfur reduction (Blumenthals et al., 1990). Two different enzymes (sulfhydrogenase and sulfide dehydrogenase) were identified to catalyze reduction of sulfur in P. furiosus. The “bifunctional” sulfhydrogenase was isolated from the cytoplasm and shown to be identical with the aforementioned hydrogenase. Sulfhydrogenase can reduce both, sulfur and polysulfide, and oxidize H2 (Ma et al., 1993). Isolation of the coding genes for sulfhydrogenase revealed that this enyzme actually consists of four subunits (b,g,d and a) encoded in a transcriptional unit, hydBCDA. Homology studies revealed a similarity of HydB and HydG with subunits of sulfite reductase from Salmonella typhimurium (Pedroni et al., 1995). Further biochemical and spectroscopic studies provided a more detailed insight into the molecular architecture of sulfhydrogenase and revealed that more [FeS] clusters were present in this enzyme than previously identified. The hydrogenase activity is localized at the adsubunits and the sulfur reductase activity at the bg-subunits. Redox centers are proposed to be arranged as follows. Three [4Fe-4S] cubanes reside in the d-subunit, two [4Fe-4S] cubanes in the b-subunit, one [2Fe-2S] cluster and one FAD in the g-subunit and the NiFe center in the asubunit (Arendsen et al., 1995; Silva et al., 1999a). Sulfide dehydrogenase, which was also identified in the cytoplasm, catalyzes the reduction of polysulfide to H2S with NADPH as electron donor. This enzyme was found to have a heterodimeric structure and to contain flavin and four [FeS] centers (Ma and Adams, 1994). A possible physiological role of sulfhydrogenase and sulfide dehydrogenase is assumed to be that of an electron sink (Ma and Adams, 1994; Hedderich et al., 1999). During fermentative degradation of glucose to acetate, liberated electrons are transferred to ferredoxin by oxidoreductases (Schäfer and Schönheit, 1992;

731

Mukund and Adams, 1991). Reoxidation of reduced ferredoxin could be directly achieved by sulfhydrogenase. Moreover, NADPH accumulating during glutamate fermentation (Robb et al., 1992) could be reoxidized by sulfide dehydrogenase. The formation of high concentrations of H2S from sulfur with H2 or methanol has been observed in cultures of methanogens (Stetter and Gaag, 1983a). However, growth due to this reaction has not been demonstrated.

Metabolism of Organic Electron Donors Carbohydrates Most of the archaeal sulfurreducers grow either lithotrophically on H2 or heterotrophically on complex substrates such as meat or yeast extracts. Only a few isolates like Thermoproteus species and Pyrococcus species were found to utilize defined carbohydrates (for review see Adams, 1994; Schönheit and Schäfer, 1995; Kengen et al., 1996; Hedderich et al., 1999). The thermoacidophilic, sulfur-reducing archaeon Thermoproteus tenax utilizes, besides other substrates, glucose for growth by sulfurrespiration (Zillig et al., 1981; Fischer et al., 1983). Part of the glucose can be transiently stored as glycogen (König et al., 1982). Glucose in the energy metabolism is completely oxidized (Selig and Schönheit, 1994). Labeling experiments with [13C]- and [14C]-glucose and enzymatic studies demonstrated that T. tenax employs in parallel a modified EmbdenMeyerhof-Parnas (EMP) pathway and the nonphosphorylated Entner-Doudoroff (ED) pathway to metabolize glucose (Siebers and Hensel, 1993; Selig et al., 1997). Of the two pathways, the EMP-pathway is used predominantly for glucose metabolism. It was suggested that the preference for one of the two pathways is regulated in response to physiological conditions (Schönheit and Schäfer, 1995). The key enzyme of the modified EMP-pathway in T. tenax is the PPi-dependent phosphofructokinase (PPi-PFK). In contrast to ATP-PFK, which is present in most organisms, the PPi-PFK uses PPi rather than ATP to phosphorylated fructose-6-phosphate. Purified PPi-PFK from T. tenax was found to be a multimeric enzyme of ca. 100 kDa mass and not to be regulated by ATP, ADP or fructose-2,6bisphosphate, the classical effectors of ATP-PFK (Siebers et al., 1998). Phylogenetic analysis of the PPi-PFK encoding gene sequence demonstrated that the T. tenax PFK is of early descent (Siebers et al., 1997). Glucose dehydrogenase, which is the first enzyme of the ED pathway, was also purified from T. tenax. The active form of the enzyme is a homodimer with a total mass of 84 kDa and uses NADP+ as cosubstrate for

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glucose oxidation (Siebers et al., 1997). Decarboxylation of pyruvate to acetyl-CoA is catalyzed by pyruvate:ferredoxin oxidoreductase (Selig and Schönheit, 1994; Schönheit and Schäfer, 1995). This enzyme is also operative in Pyrobaculum islandicum and Pyrococcus furiosus (Schäfer and Schönheit, 1991). Further oxidation of acetate to CO2 in T. tenax and P. islandicum involves the citric acid cycle (Selig and Schönheit, 1994). The sugar metabolism in Pyrococcus furiosus, which has been intensively investigated, has many parallels to that in Thermoproteus tenax (Mukund and Adams, 1991; Schäfer and Schönheit, 1992; Kengen et al., 1994; Kengen et al., 1996; Schäfer et al., 1994). P. furiosus, in which sulfur reduction facilitates fermentation, has no capability for complete oxidation and forms acetate as an organic end product (Schönheit and Schäfer, 1995). Peptides Thermoproteus tenax and Pyrobaculum islandicum (Huber et al., 1987) have been reported to utilize peptides with sulfur as electron acceptor. Considering the capacity of these two archaea to oxidize acetate completely to CO2 via the citric acid cycle (Selig and Schönheit, 1994), it can be assumed that also peptides are completely oxidized to CO2. Autotrophic Carbon Assimilation Thermoproteus neutrophilus (Zillig et al., 1981; Fischer et al., 1983) is a facultative autotroph that can use either CO2 or acetate as carbon source during growth on H2 and sulfur. The pathway for CO2 fixation was studied by 14C-labeling experiments and measurement of enzyme activities. The key enzyme of the Calvin-cycle, ribulose1,5-bisphosphate carboxylase (for summary see Watson and Tabita, 1997), was not detected in extracts of T. neutrophilus cells. Results rather suggested the presence of a reductive citric acid cycle (Schäfer et al., 1986). Enzyme activities corroborating this CO2 fixation pathway, including the ATP citrate lyase, were subsequently demonstrated (Beh et al., 1993). Acidianus is a genus of facultatively anaerobic Archaea that can grow aerobically by sulfur oxidation or anaerobically by sulfur reduction with H2. In both cases, growth is autotrophic (Segerer et al., 1986; Segerer and Stetter, 1992). Enzyme studies with extracts of autotrophically grown A. infernus cells indicated that acetylCoA carboxylase and propionyl-CoA carboxylase function as the main CO2-fixation enzymes. A 3-hydroxypropionate cycle is proposed for this organisms as route of CO2 fixation (Menendez et al., 1999), where two moieties of CO2 are fixed

CHAPTER 1.22

by the aforementioned enzymes, and glyoxylate is formed for further synthesis of organic compounds from malyl-CoA, while acetyl-CoA is concomitantly regenerated. This pathway has originally been detected in the phototrophic bacterium Chloroflexus aurantiacus (Holo, 1989; Straußet al., 1992). Detoxification of Superoxide Superoxide reductase (SOR), was purified from Pyrococcus furiosus, and proposed to function in scavenging superoxide via a net reduction to H2O2 rather than via dismutation to H2O2 and O2 as is known from superoxide dismutase (Jenney et al., 1999). Reduced rubredoxin is suggested as the primary source of reducing power for SOR. Reduction of rubredoxin is catalyzed by NAD(P)H:rubredoxin oxidoreductase that has also been purified from P. furiosus (Ma and Adams, 1999). Reductive scavenging of superoxide appears to be a widespread mechanism in anaerobes to protect against the deleterious superoxide species. Homologs of the SOR encoding gene have been identified in many complete genomes of anaerobic microorganisms, but not in those of aerobic organisms. In contrast, genes coding for superoxide dismutase are not generally present in anaerobic microorganisms. The SOR encoding gene also shows homology to the redox proteins desulfoferredoxin and neelaredoxin from Desulfovibrio desulfuricans and D. gigas, respectively (Jenney et al., 1999).

Microorganisms Reducing Sulfur Compounds Other Than Sulfate or Sulfur Bacteria The capacity for the dissimilatory reduction of sulfur compounds other than sulfate and sulfur, especially sulfite and thiosulfite, has been frequently observed among sulfatereducing microorganisms and also among some sulfur-reducing microorganisms. However, there are also prokaryotes that neither reduce sulfate nor elemental sulfur but instead utilize other sulfur compounds as electron acceptors. There are several reports on tetrathionate and thiosulfate reduction in bacteria other than sulfate or sulfur reducers (Barrett and Clark, 1987). A reduction of these sulfur species seems to be abundant especially among enterobacteria. Often, the capacities to reduce tetrathionate and thiosulfate coincide, which is explained by one reductase for both compounds (Oltmann et al., 1975; Barrett and Clark, 1987). Tetrathionate is first reduced to thiosulfate. In Citrobacter and Proteus, there is evidence for an electron transport chain to tetrathionate allowing respiratory

CHAPTER 1.22

Dissimilatory Sulfate- and Sulfur-Reducing Prokaryotes

energy conservation (Oltmann et al., 1975; Novotny and Kapralek, 1979); the growth substrates were sugars. A complete oxidation (viz. acetyl-CoA oxidation) associated with tetrathionate reduction has not been reported for the enterobacteria. Formed thiosulfate may be further reduced to sulfide and sulfite, the latter being often an end product that is not reduced further. For energetic reasons, such an incomplete reduction of thiosulfate probably does not allow a chemiosmotic process and thus appears to be a by-reaction. Suspensions of the phototroph, Thiocapsa floridana, reduced thiosulfate with endogeneous hydrogen donors in the dark (Trüper and Pfennig, 1966). A marine Pseudomonas-like strain grew anaerobically on lactate in the presence of thiosulfate or sulfite and formed sulfide. Lactate alone was not utilized (Tuttle and Jannasch, 1973). It is unknown whether sulfite reduction was of a respiratory type or just a facilitated fermentation. Reduction of thiosulfate or sulfite, probably as a mere hydrogen sink, has also been observed in mesophilic and thermophilic saccharolytic clostridia. The sulfite reductase in Clostridium pasteurianum was induced by sulfite and distinctive from the assimilatory enzyme (Harrison et al., 1984). Even yeast cells catalyzed a reduction of thiosulfate to sulfite and sulfide, and of sulfite to sulfide (Neuberg and Welde, 1914; Hollaus and Sleytr, 1972; McCready and Kaplan, 1974; Stratford and Rose, 1985); sulfite reduction was observed in an aerated culture. The reductions seemed to be by-reactions. An organic sulfur compound used by several bacteria as electron acceptor is (CH3)2SO, dimethylsulfoxide (DMSO), which is reduced to dimethylsulfide (DMS). The utilization of DMSO was first shown in the phototroph Rhodobacter capsulatus that did not grow anaerobically on sugars in the dark unless the acceptor was added (Yen and Marrs, 1977). It was first assumed that DMSO serves merely as a H2 sink allowing substrate-level phosphorylation (Madigan and Gest, 1978; Madigan et al., 1980). Later, however, Rhodobacter capsulatus and also Rhodospirillum rubrum were reported to grow by DMSO reduction most likely in a respiratory manner (Schultz and Weaver, 1982). Among other substrates, also acetate allowed anaerobic growth in the dark when DMSO was present. Earlier, a definitive respiratory DMSO reduction had been already shown with a spirillum isolated with lactate as electron donor (Zinder and Brock, 1978a). The spirillum grew with H2 if acetate as a carbon source and some yeast extract as sulfur source for assimilation were present. In addition to DMSO, the organism reduced sulfur, sulfite and thiosulfate and resembled Desulfos-

733

pirillum delyianum (spirillum 5175) isolated with sulfur (Wolfe and Pfennig, 1977). The latter also turned out to reduce DMSO (N. Pfennig, personal communication). Anaerobic growth due to DMSO reduction with H2 was also observed with Escherichia coli (Yamamoto and Ishimoto, 1978). The obvious respiratory character of DMSO reduction was confirmed by measurements with H2 and glycerol (Bilous and Weiner, 1985). However, as with nitrate (Thauer, 1988), there is no evidence that DMSO is an electron acceptor for acetyl-CoA oxidation in E. coli; the citrate cycle is not operative in E. coli under anoxic conditions, and acetate is an end product. Some other enterobacteria, Pseudomonas aeruginosa and Bacillus subtilis, reduced DMSO in complex glucose medium (Zinder and Brock, 1978b). However, DMSO reduction seemed to be not very effective (90%) and are functionally redundant. They have significant similarity with NorB of other nitric oxide reductases such as that of P. stutzeri, with the exception of an N-terminal extension of about 300 residues. These extra residues of the R. eutropha enzymes likely add two additional membrane-spanning regions and a large hydrophilic loop. The primary sequence of the N-terminal extension does not have similarity with other known proteins. Part of its function may relate to the observation that there is no evidence for a gene encoding a NorC equivalent in R. eutropha. This suggests that the amino terminal extension is functionally equivalent to NorC, but no direct evidence has been provided in support of this conclusion. A gene whose product is similar to the NorB and NorZ products has turned up in the genome of Synechocystis strain PCC 6803 (Kaneko, 1996). Interestingly, genes encoding nitrite and nitrous oxide reductase were not identified in this bacterium. Because PCC 6803 can not reduce nitrite, it has never been considered as capable of gaseous nitrogen oxide metabolism. This makes the presence of nitric oxide reductase unexpected. However, as discussed above, other strains have recently been characterized which have nitric oxide reductase but lack nitrite reductase, suggesting this mode of truncation of the denitrification electron transport chain may be fairly prevalent. Ongoing genome sequencing efforts in N. meningitidis (Sanger Centre, 1999) and N. gonorrhoeae (Roe, 1999) have also identified genes whose products are similar to the R. eutropha nitric oxide reductases. Neisseria and R. eutropoha are both in the b-subgroup of the proteobacteria (Table 1), but it is unclear if this class of enzymes will be preferentially found in one taxonomic group. Alignment of nitric oxide reductases also demonstrates that the R. eutropoha-type nitric oxide reductases have other unique sequence motifs not found in the P. stutzeri type reductases (Fig. 4). The reduction of nitric oxide to nitrous oxide occurs at the binuclear center. Current models suggest that two nitric oxide molecules bind at the active site, although it is not clear if nitric oxide binds to both metal centers or if a dinitrosyl complex is formed (Ye, 1994; MoenneLoccoz, 1998). Electrons enter the Nor complex through the cytochrome c in NorC and then flow to the six-coordinate heme in NorB. Electrons are then transferred to the binuclear center. The high affinity of reduced heme for nitric oxide

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makes it unclear if reduction of the catalytic site involves formation of a ferrous heme-nitric oxide species. Irrespective of the exact electron transfer steps, it is clear that proximity of the two nitric oxide molecules is critical in the formation of the N-N bond.

Nitrous Oxide Reductase The presence of nitrous oxide reductase can be identified independently of the other reductases because many organisms can grow with nitrous oxide as sole terminal oxidant. This has led to the identification of nitrous oxide reductase in several bacteria that are not complete denitrifiers (Yoshinari, 1980; Payne, 1982). The purification and characterization of nitrous oxide reductase was difficult because its activity is lost in cell extracts. Nutritional studies had identified copper as an essential nutrient for nitrous oxide reductase activity (Matsubara, 1982) and further work led to the isolation of a soluble copper protein which, under the proper conditions, had nitrous oxide reductase activity (Zumft, 1982). Additional studies have demonstrated that enzyme purified anaerobically and assayed using reduced viologens as electron donor has the highest specific activity. The latter observation is somewhat puzzling because the natural electron donors to nitrous oxide reductase are likely to have much higher redox potential. Nitrous oxide reductases from several complete denitrifiers have been extensively characterized (Riester, 1989; SooHoo, 1991; Hulse, 1990; Snyder, 1987). These enzymes are all related and are multi-copper periplasmic proteins. The protein appears to be homodimeric in most preparations, with four coppers per subunit. Current data suggest there are two binuclear copper centers in the active enzyme. These centers undergo spectroscopic shifts depending on their redox state, and this has resulted in the characterization of different colored forms of the enzyme. One of the copper centers has been structurally defined as a CuA site. The CuA center was described originally in the heme copper oxidases. However, the exact nature of this site was unclear until the related site in nitrous oxide reductase was characterized. Analysis of the deduced primary sequence of the nitrous oxide reductase identified a structural motif found otherwise only in the CuA-containing subunit of cytochrome oxidase (Viebrock, 1988). Additional spectroscopic studies provided further evidence of the similarities between one of the sites in nitrous oxide reductase and the CuA site of cytochrome oxidase (Farrar, 1991; Scott, 1989; Antholine, 1992). A more precise understanding of the structure of the CuA center has been

CHAPTER 1.23

obtained with the determination of the highresolution structure of two cytochrome oxidases (Tsukihara, 1995; Ostermeier, 1997). In both cytochrome oxidase and nitrous oxide reductase, the CuA site has a role in transferring electrons from external electron donors to the active site. The other copper center in nitrous oxide reductase is, by exclusion, assumed to be the site of nitrous oxide binding and reduction. This site has been designated CuZ. Initial studies suggested both CuZ and CuA sites were bis-thiolatebridged dinuclear copper sites (Farrar, 1991). Isolation of spectral signals arising from the CuZ center has proved difficult. Spectroscopic analysis of the CuZ signals has relied on poising the enzyme in specific oxidation states to make the CuA center Electron Paramagnetic Resonance (EPR) or optically silent. However, it has been suggested that the signals assigned to the CuZ site are actually different redox states of the CuA center (Farrar, 1998). In this latter work, it was shown that both the CuA site and the putative CuZ site were bis-thiolate-bridged dinuclear copper sites. This would require, if these were separate sites, four conserved Cys residues for formation of the two binuclear centers. Alignment of deduced nitrous oxide reductase sequences does not identify four conserved Cys residues. Instead, there are only two absolutely conserved Cys residues in nitrous oxide reductase. It is possible that the position of a Cys is shifted in one sequence relative to others accounting for the deficiency of conserved residues. However, mutation of one of the Cys residues outside the CuA domain does not lead to a loss of nitrous oxide reductase activity (Dreusch, 1996). Taken together, these data are not consistent with assuming that there are two bisthiolate-bridged dinuclear copper sites in nitrous oxide reductase. If conserved residues do not bridge the catalytic center, what then is its structure? Alignment of available nitrous oxide sequences does indicate there are a number of completely conserved His residues. The catalytic center might therefore be a His-ligated structure similar to the Type 3 copper centers found in enzymes such as laccase (Solomon, 1996). However, there is currently no direct evidence for the presence of such a center. Given the uncertainty in the structure of the catalytic center, it is difficult to develop a useful model of the nitrous oxide reductase catalytic cycle. Nitrous oxide is chemically inert and also a poor ligand, making its reduction an interesting problem in transition metal-ligand chemistry (Kroneck, 1990). Other enzymes, including nitrogenase (Jensen, 1986) and carbon monoxide dehydrogenase (Lu, 1991), also have been found to have nitrous oxide reductase activity, indicat-

CHAPTER 1.23

ing that it is possible that transition metals other than copper can reduce nitrous oxide. Only copper-containing nitrous oxide reductases have been purified from complete denitrifiers. However, there appear to be variants of this structure present in other bacteria. The nitrous oxide reductase from W. succinogenes has been purified and shown to contain both copper and iron (Teraguchi, 1989; Zhang, 1991). The iron is attributed to an associated cytochrome c, though EPR suggests the presence of a CuA site. Because the primary structure of the W. succinogenes enzyme is unknown, the relationship of this enzyme to other nitrous oxide reductases is unknown. Another novel nitrous oxide reductase has been suggested to occur in Flexibacter canadensis (Jones, 1990). Most nitrous oxide reductases are inhibited by acetylene (Balderston, 1976; Yoshinari, 1976). However, the nitrous reductase from F. canadensis is insensitive to this compound (Jones, 1990). Preliminary characterization of this enzyme suggests it is membrane-associated, further differentiating it from other reductases (Jones, 1992). The nitrous oxide reductase from R. sphaeroides IL106 was also thought to be divergent because it was reported to contain Zn and Ni in addition to copper (Michalski, 1986). However, a more rigorous characterization of this enzyme suggests it is similar to typical copper-containing enzymes (Sato, 1998).

Genetics of Denitrification Gene Organization Analysis of the structure and organization of denitrification genes has been investigated in denitrifiers that are primarily members of the proteobacteria. The most extensive characterizations have been carried out with genomes of the pseudomonads as well as that of P. denitrificans strains. Although no true denitrifier has had its chromosome sequenced, there are several projects to sequence organisms that are partial or complete denitrifiers. Nitrite and Nitric Oxide Reductase The most extensive examinations of denitrification gene organization have involved P. aeruginosa and P. stutzeri in which the nitrite reductase and nitric oxide reductase structural genes are about 10 kb apart (Arai, 1995; Braun, 1992). This tight linkage is found also in P. denitrificans (Baker, 1998). In general, such tight linkage of the nir and nor gene clusters is not observed in those denitrifiers containing a copper-type nitrite reductase. In R. sphaeroides strains IL106 and 2.4.3, the genes for nitrite reductase are not closely linked to the

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genes for nitric oxide reductase (Schwintner, 1998; Tosques, 1997). This is also the case in N. gonorrhoeae (Roe, 1999) and N. meningitidis (Sanger Centre, 1999). One exception is Pseudomonas G-179 (which is apparently a member of the a proteobacteria subclass) where it has been shown that the gene encoding the copper-type nitrite reductase is in a cluster with the genes for nitric oxide reductase and the periplasmic nitrate reductase (Bedzyk, 1999). Genes encoding proteins required for assembly of a particular reductase are typically found clustered with the structural gene for that particular reductase. For example, in those bacteria encoding a cd1-type nitrite reductase, several genes whose products are involved in the synthesis of d1 heme will be required for the production of an active enzyme. Systematic inactivation of genes closely linked to the nitrite reductase structural gene has identified a number of genes whose products are involved in heme d1 biosynthesis (Kawasaki, 1997; Palmedo, 1995). Although this has led to the identification of a set of genes uniquely required for heme synthesis during denitrification, the details of the biosynthesis of heme d1 are not elucidated as yet. The organization of the genes for assembly of the cd1 protein is dissimilar in every denitrifier. In P. aeruginosa, eleven adjacent genes have been identified whose products are suggested to be involved in nitrite reductase activity (Arai, 1994). These genes are transcribed in the same direction and are postulated to be transcribed in a single transcript (Kawasaki, 1997). In P. stutzeri, this cluster of genes has been rearranged so that the genes are no longer adjacent and at least three different transcripts are produced (Palmedo, 1995). The total number of proteins that is required for expression of an active copper-containing nitrite reductase would be expected to be less than with the heme-type nitrite reductase. Examination of sequence flanking the structural gene encoding the copper-type nitrite reductase in N. gonorrhoeae (Roe, 1999), N. meningitidis (Sanger Centre, 1999), R. sphaeroides 2.4.3 and Pseudomonas G-179 (Bedzyk, 1999) revealed only one other conserved gene. In R. sphaeroides 2.4.3, this undesignated gene is located about 200 bp downstream of the putative translation stop of nirK (the nitrite reductase structural gene). Preliminary evidence suggests it is transcribed from the nirK transcription start. The role of the product of this gene is unclear. The difference in the amount of DNA required to produce an active copper-containing nitrite reductase versus the heme-containing nitrite reductase is notable. In R. sphaeroides 2.4.3, about 2.5 kb are required to encode the copper-

780

J.P. Shapleigh

containing nitrite reductase and the accompanying gene of unknown function. In P. aeruginosa, about 10 kb appear to be required to code for proteins necessary for assembly of an active nitrite reductase. The structural genes for the heterodimeric form of nitric oxide reductase, designated norC and norB, have been sequenced from a number of denitrifiers. In every case, the transcription start precedes norC and the gene order is norCB. The complete nor operon consists of norCB and one or two additional genes. For example, in P. denitrificans and R. sphaeroides norQ and norD follow norB (Bartnikas, 1997; De Boer, 1996). In P. aeruginosa and P. stutzeri, norD follows norB (Arai, 1995; Zumft, 1994). In both these pseudomonads norQ (designated nirQ) is present, but it is immediately upstream, and divergently transcribed from the structural gene for nitrite reductase, nirS (Arai, 1994; Jungst, 1992). Inactivation of norQ or norD leads to a loss of nitric oxide reductase activity but does not appear to inhibit assembly of nitric oxide reductase (Jungst, 1992; Mitchell, 1998; De Boer, 1996). This suggests the likelihood that both NorQ and NorD are accessory proteins required for the assembly of nitric oxide reductase. One possible role would be insertion of non-heme iron. Neither norQ nor norD has been found in those denitrifiers encoding the single subunit type of nitric oxide reductase. There are no obvious orthologs of either norQ or norD present in the chromosome of Synechocystis sp. strain PCC 6803 (Kaneko, 1996). Nor have norQ or norD orthologs been identified in the ongoing genome sequencing efforts in N. gonorrhoeae or N. meningitidis. The absence of these proteins in these denitrifiers is somewhat surprising given the sequence similarity of the single subunit and heterodimeric nitric oxide reductases. The sequence of the R. eutropha norB is preceded by an open reading frame (ORF) encoding a protein containing a high percentage of His residues (Cramm, 1997). It is possible the product of this gene might play a role in assembly of an active nitric oxide reductase. R. eutropha is unusual in that it contains two nitric oxide reductase structural genes (Cramm, 1997). One of these, norB, is located on a megaplasmid while the other, norZ, is located on the chromosome. This is the only bacterium described in which functionally redundant terminal N-oxide oxidoreductases have been observed. It is also interesting that in R. eutropha norZ does not appear to be tightly linked to the nitrite reductase structural gene. R. eutropha contains a cd1-type nitrite reductase. This genetic organization is different from those bacteria containing a heterodimeric nitrite oxide reductase and a cd1-type nitrite reductase.

CHAPTER 1.23

In P. denitrificans, there are two additional genes, designated norE and norF, whose products appear to be required for nitric oxide reductase activity (De Boer, 1996). These genes are located immediately downstream of the norCBQD cluster. Insertional inactivation of either norE or norF reduces nitric oxide reductase activity but does not significantly affect its expression. Orthologs of norE have been found in other denitrifiers, for example, the ORF175 protein in P. stutzeri (Glockner, 1996) and in Pseudomonas sp. G-179 (Bedzyk, 1999). However, no norE ortholog has been identified in R. sphaeroides or in those bacteria encoding the single subunit nitric oxide reductase. No obvious norF orthologs have been identified, but genes whose products have some similarity to norF have been described in P. stutzeri (ORF82) and in Pseudomonas sp. G-179 (Bedzyk, 1999). The role of norE and norF remain undetermined. However, because of its similarity to the subunit III of cytochrome oxidases, it has been suggested NorE is a third subunit of the nitric oxide reductase protein complex (De Boer, 1996). Experimental conformation for NorE as a part of an active nitric oxide reductase complex is lacking. Nitrous Oxide Reductase While the nir and nor gene clusters are often linked, the location of the nos gene cluster (nos refers to genes related to nitrous oxide reductase and not to genes for nitric oxide synthase) relative to other denitrification genes is more variable. In P. stutzeri, the genes encoding nitrous oxide, nitrite and nitric oxide reductase are within a stretch of about 30 kb (Jungst, 1991). In P. aeruginosa, the nitrous oxide reductase gene cluster is not as tightly linked to the nitrite and nitric oxide reductase genes (Vollack, 1998). However, the genes for all four nitrogen oxide reductases are located within the 20- to 36-min segment of the P. aeruginosa chromosome. In several denitrifiers, including R. eutropha (Zumft, 1992), Sinorhizobium meliloti (Holloway, 1996) and R. capsulatus (Rhodobacter, 1999), nos genes are found on plasmids. By comparison, all denitrifiers characterized in detail have both nir and nor located on the chromosome. The variable location of the nos cluster may reflect nitrous oxide’s lack of toxicity, and therefore the accumulation of nitrous oxide that would follow the loss of nitrous oxide reductase activity has only limited consequences. Gene organization within the nos cluster is much more conserved than is that in the nir or nor clusters. In almost every sequence, nosR is immediately upstream of nosZ, and nosZ is typically followed by nosDFYL. Undefined genes clustered with nosZ are likely to be involved in assembly of an active nitrous oxide reductase,

CHAPTER 1.23

perhaps in copper incorporation. Inactivation of nosF, nosD or nosY leads to production of an inactive nitrous oxide reductase (Zumft, 1990). Sequence analysis indicates that nosY encodes a membrane-bound protein, nosD a periplasmic protein, and nosF a cytoplasmic nucleotidebinding protein. It has been suggested these proteins form a complex involved in copper processing and insertion into nitrous oxide reductase (Zumft, 1997). In S. meliloti, a gene that has been designated nosX follows nosL. Inactivation of nosX causes a loss of nitrous oxide reductase activity in S. meliloti (Chan, 1997). Possible nosX orthologs have been found in other denitrifiers including “A. cycloclastes” (McGuirl, 1998) and B. japonicum (Genbank accession number {AJ002531}). A gene encoding a product similar to the nosX product has also been identified in P. denitrificans, but since it is part of the nir gene cluster it has been designated nirX (Genbank accession number AJ001308). A nosX ortholog has not been identified as yet in the peudomonads. The role of nosX is unclear but current data do not suggest a role in copper processing. Isolation of mutants of P. stutzeri unable to use nitrous oxide as sole terminal oxidant led to the isolation of an additional gene whose product is required for nitrous oxide reductase activity. This gene was designated nosA, and it was shown to encode an outer membrane protein that is required for copper transport into the cell (Lee, 1991; Lee, 1989). Inactivation of nosA results in expression of the nitrous oxide reductase apoprotein. Putative nosA orthologs have been found in other denitrifiers, including P. aeruginosa (Yoneyama, 1996), but nosA is not found in the nos cluster of P. stutzeri. This organization may be because the nosA product is playing a more general role in cell physiology.

Nitrate Reductase Of the various nitrate reductases, the respiratory and periplasmic forms have been studied in the most detail in the context of denitrification. Genes encoding the respiratory nitrate reductase, designated nar, have been completely sequenced in P. aeruginosa (Genbank accession number {Y15252}) and partially sequenced in P. denitrificans (Berks, 1995) and Pseudomonas fluorescens (Philippot, 1997). The nar genes are not clustered with other genes required for denitrification in P. aeruginosa (Vollack, 1998). The relatively limited interest in the nar genes in denitrifiers is due to the extensive study of nar genes in E. coli and other non-denitrifiers. Genes encoding the periplasmic nitrate reductase have been characterized in several denitrifi-

The Denitrifying Prokaryotes

781

ers including P. denitrificans GB17 (Berks, 1995) and R. sphaeroides (Reyes, 1996). The structural genes for this nitrate reductase are napA and napB: napA encodes the molybdopterin cofactor and napB encodes the cytochrome c containing subunit. These two genes are clustered with other genes that have been designated napCDE. The napC gene encodes a membrane-bound cytochrome c, which is the likely electron donor for the periplasmic nitrate reductase; napD encodes a cytoplasmic protein and napE a small membrane protein. The function of these genes’ products is unknown. The nap genes of P. aeruginosa are present on the chromosome but are not tightly linked to nar or other denitrificationrelated gene clusters (Vollack, 1998). In R. eutropha (Siddiqui, 1993)and R. sphaeroides strain 2.4.1 (Schwintner, 1998), the nap genes are localized on plasmids. In R. sphaeroides IL106, the nap genes are apparently located on the chromosome (Schwintner, 1998). In Pseudomonas G-179, the nap genes are part of a large cluster which includes nir and nor genes (Bedzyk, 1999).

Additional Genes Required for Denitrification A few other genes frequently associated with nitrogen oxide gene clusters deserve mention. One is hemN, which encodes an oxygenindependent coproporphyrinogen oxidase (Gibson, 1992). R. sphaeroides encodes two hemN paralogs, the second of which is designated hemZ (Zeilstra-Ryalls, 1995). The R. sphaeroides hemN is clustered with the genes encoding nitric oxide reductase (unpublished). Inactivation of hemN (which was originally designated hemF) inhibits the ability to grow anaerobically under any conditions tested (Gibson, 1992). Aerobic growth is not affected because there is an oxygen-dependent form of this enzyme. The role of the hemZ product is unclear. Interestingly, hemZ is adjacent to fnrL whose product is an important regulator of anaerobically expressed genes (Zeilstra-Ryalls, 1995). These two genes are clustered with the ccoN operon that encodes the cbb3-type oxidase. The cbb3-type oxidase is a heme-copper enzyme and is the oxidase with the highest level of similarity to nitric oxide reductase (Saraste, 1994). It is notable then that both hemZ and hemN are clustered with related terminal oxidoreductases and regulatory proteins important in maintaining anaerobic physiology. It is possible that this gene arrangement may have occurred by gene duplication providing further evidence for the evolutionary link between aerobic respiration and denitrification.

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J.P. Shapleigh

The hemN gene is clustered with denitrification genes in Pseudomonas sp. G-179 (Bedzyk, 1999) and is clustered with the ccoN genes in both P. denitrificans (van Spanning, 1997) and P. aeruginosa (Zumft, 1997). A hemN ortholog has not been found in the nor cluster in P. denitrificans or P. aeruginosa. The regulation of hemN in P. aeruginosa (discussed in more detail below) further emphasizes the importance of the activity of the hemN product during denitrification (Rompf, 1998). Sequencing of the nor cluster in R. sphaeroides (Bartnikas, 1997) and nos region in P. stutzeri (Glockner, 1996) has revealed a gene present in both regions whose product may be important to the physiology of denitrification. This gene has been designated nnrS in R. sphaeroides and orf396 in P. stutzeri (Fig. 5). The product of the genes from both bacteria is a membrane protein probably containing twelve membrane-spanning regions. In R. sphaeroides, nnrS is expressed only during denitrification and is regulated by NnrR, which also regulates nirK and nor (unpublished). Inactivation of nnrS has no obvious effect on growth under any conditions (unpublished). Though nnrS orthologs have not been found in the denitrification gene clusters of other well studied denitrifiers, examination of ongoing genomic sequencing efforts reveal the presence of nnrS orthologs in every denitrifier. In R. capsulatus, a nnrS ortholog is present on a plasmid and is closely linked to the nos genes (Rhodobacter, 1999). There is also an nnrS ortholog located on the chromosome. Copies of genes encoding products similar to nnrS have been identified in the N. gonorrhoeae (Roe, 1999), N. meningitidis (Sanger Centre, 1999) and P. aeruginosa chromosome (Pseudomonas, 1999), although none of these are clustered with denitrification genes. Significantly, nnrS orthologs have not been found in non-denitrifying bacteria. The function of nnrS has not been determined, but it is obviously not essential for denitrification or it would likely have been isolated in mutant screens. However, work in R. sphaeroides and its distribution among the bacteria indicates nnrS is a denitrificationassociated gene. It seems likely that as more work is done on the genetics of denitrification, many genes that are not essential to, but whose products are physiologically important for, denitrification will be described.

Regulation of Genes Required for Denitrification This section will focus primarily on the regulation of those genes encoding respiratory nitrogen oxide reductases and genes required for their

CHAPTER 1.23

assembly. As denitrification depends on the presence of nitrogen oxides, it is natural to describe denitrification genes as part of a stimulon, a term that refers to operons responding together to a particular environmental stimulus (Neidhardt, 1990). In general, the regulators of denitrification can be differentiated into the nitrate, nitric oxide, and nitrous oxide stimulons. The organization of denitrification genes roughly reflects the organization of the stimulons. The nitrate and nitrous oxide stimulons are primarily made up of the nar and nos gene clusters, respectively. These two, independently regulated, gene clusters are not linked to each other and are frequently distant from other denitrification-related clusters. The nitric oxide stimulon is made up of both the nir and nor clusters. These two gene clusters are the most strongly linked of any of the denitrification gene clusters. As denitrification is, in most cases, an anaerobic process, other stimulons and regulons required for anaerobic growth overlap the denitrification-related stimulons. This overlap can make it difficult to differentiate regulatory factors that directly modulate gene expression from those that indirectly affect gene expression. This discussion will focus on those proteins that current data suggest are directly involved in regulation of the nitrogen oxide reductases. It is important to note that a particular stimulon will likely include genes whose products are not directly required for denitrification and, consequently, not covered in this review. However, this does not minimize the usefulness of organizing denitrification genes into stimulons. One other important consideration is the relationship of denitrification and oxygen respiration. Denitrification is primarily an alternative form of respiration inasmuch as oxygen is a preferred oxidant. However, this does not imply that all denitrifiers have the same set point at which oxygen respiration is switched to nitrogen oxide respiration. Available data suggest that the onset of expression of denitrification genes occurs over a wide range of oxygen concentrations. The best example of an aerobic denitrifier is P. denitrificans GB17, which was originally described as an aerobic denitrifier (Robertson, 1984). In addition, other strains have been described as aerobic denitrifiers (Bonin, 1991; Patureau, 1994; van Niel, 1992; Frette, 1997; Robertson, 1995; Ka-Jong, 1997). None of the intensively studied model denitrifiers activate gene expression, even at moderate concentrations of oxygen. Therefore, the molecular mechanisms that permit aerobic denitrification are not currently understood. Nitrate Stimulon The presence of nitrate and a reduction in oxygen tension stimulates

CHAPTER 1.23

The Denitrifying Prokaryotes

783

NmNnrS NgNnrS PsORF396 PaNnrS RsNnrS RcNnrS

------------------- MKFT-KHPVWAMA ---FRPFYSLAALYGALSVLLWGFGYT------------------- MKFT-KHPVWAMA ---FRPFYSLAALYGALSVLLWGFGYT------------ MQUIDRRKALS-IAPIWRLA ---FRPFFLAGSLYALLAIPBWVAWWTG --------------------- MA-IPPIWRLG ---FRPFFLGGALFAVLAIALWLAALAG ------------- MASDRPRTYT-GPALLSYG ---FRPFFLLSALFAAGAUPUWLAUWSMAHPHLDSPAAGRHHPDIPAAGAPMTALLRLLSDSFRUFFLLASLWAAAAMALWLWWLWQ ** *: * :*

NmNnrs NGNNRS PsORF396 PsNnrS RcNnrS RcNnrS

--- G-THELSGFY ----WHAHEMIWGYAGLUUIAFLLTAUATWTGBGHQPPTRGGULTIF --- G-THELSGFY ----WHAHEIWGYAGLUUIAFLLTAUATWTGQPPPTRGGULUGLTAF LWPG-FQPTGGWLA ---WHRHEMLFGFAMAIUAGFLLTAUQTWTGQTAPSGNRLUGLAAU LWSG-FQPTGGWLA ---WHRHEMLFGFGUAIIAGFLLTAUQTWTGUPGLQGRPLALLAGL --- G-RIGLAGPFSPIDWHIHEMLFGYTSAUIAGFLFTAIPNWTGRMPRRGLPLAALAAL NQIGPGGDLPNALAPSHWHAHELIFGFGMAATAGFFLTAAPNWTGKPUAGPRFIALMAGL * * * **: :: *: :: : *:: **: * * * :: : :

NmNnrs NGNNRS PsORF396 PsNnrS RcNnrS RcNnrS

WLAARIAA-FIPGWGASASGILGTLFFWYGAUCMALPUIRSQNQRNYUAUFALFULGGTH WLAARIAA-FIPGWGAAASGILGTLFFWYGAUCMALPUIRSQNRRNYUBAFAIFULGGTH WLAARLG--WLFGLPAAWLAPLDLLFLUALUWMMAQMLWAUPQKRNYPIUUULSLMLGAD WLAARLA--WLFDAPLALLLULQLSFLPLLAWAIGRSLWRURQKRNYPUUGLLLLLTLAD WIAGRFAUAGAFGTDPLLULUIDAGFLLAUTLMAUIEIAAGKNWKNLMUUGPUGLYLAAN WLAGRGAULLWGSUPPULAAGUULAFPALLTERMARQLIRRPASSEGLYLALLGLITLAE : : : : *: * * * :

NmNnrs NGNNRS PsORF396 PsNnrS RcNnrS RcNnrS

AAFHUQLHNGNLGGLLSGLQSGLUMUSGFIIGLIGTRISFFTSKRLN -----UPQIPSPK AAFHUQLHNGNLGGLLSGLQSGLUMUSGFIGLIGMRIISFFTSFFLN -----UPQIPSPK ULILTGLLQGNDALQRQGULAGLWLUAALMALIGGRUIPFFTQRGLGK ----UDAUKPWU ALULLGLFEGNDDWQRRASIAALWLIAGMMNLIGGRUIPFFTQRGLGR ----QQQUPAIA ULFHLEAMQ--QGESDIGRPLGFATUTFLIMLIGGRIIPSFTRNWLAKG --GPGPLPUPF ARULLDWLDLPPGDAAAGLRGGLAALUALUAULGGRITPQFTRNALARAGAPPAALPRSF : :: :: * *: - ** : : * :

NmNnrs NGNNRS PsORF396 PsNnrS RcNnrS RcNnrS

-WUAQASLWLPMLTAMLMAHGU----MPWLSAAFAFAAGUIFTUQUYRWWYKPULKEPML -WUAQASLWLPMLTAILMAHGU----MPWLSAAFAFAAUUIFTVQVYRWWYKPVLKEPML -WLDUALLUGTGUIALLHAFGUAMRPQPLLGLLFU-AIGVGHLLRLMRWYDKGIWKVGLL -WLDNGILLGCULUALLTAAGUTTQPTPWLAGLFA-ALGGAQLWRLWRWRDRILWQVPLL -GRFDGASLLUAUGALLCUTLA---PDAILTAALLALAAALHUURLURWRGHLUWRSPLL PWLDRSVAGCACLAALAAUFPL----SGALAGAAALALGAGQLARMGFWRSRKVLGNPLL : *:: : * : :: :: : * : :*

NmNnrs NGNNRS PsORF396 PsNnrS RcNnrS RcNnrS

WILFAGYLFTGLGLIAUGASYFKPAFLN-LGUHLIGUGGIGULTLGMMARTALGHTGNPI WILFAGYLFTGLGLIAUGASYFKPAFLN-LGUHLIGUGGIGULTLGMMARTALGHTGNSI WSLHUAMLWLUUAAFGLALWHFGLLAQSSPSLHALSUGSMSGLILAMIARUTLGHTGRPL WSLHLAYFWIAUAPLGMALWSLGLALAPSQSLHALAUGGMGGLILAMLARUTLGHTGRAL LMMHUAYGFLPLGLAATAAAAUGWASAP-AGLHLLGIGAIGGMTLAUMMRASLGHTGRPL AALHLAMLGTGIGALLQGFAAFGRGEEI-GALHVSAGUGGMILAUMADSRATLAQTGRAL : * : : : :* : : : * :: * * : * * :

NmNnrs NGNNRS PsORF396 PsNnrS RcNnrS RcNnrS

UPPPKAUPUAFWLMMAATAURMUARFSSGTAYTH---SIRTSSVLFALALLVYAWKYIPW YPPPKAUPUAFWLMMAATAURMUAUFSSGTAYTH---SIRTSSULFALALLUYAWKYIPW QLP-AGIIGAFULFNLGTAAR---UFLSUAWPUG---GLWLAAVCWTLAFALYUWRYAPM QPP-AAMPWAFALLNLGCAAR---UFLPSLLPANW--ALPLAGGLWALAFLLFAWFYAPM EAG-RALSAGFACIUAAAAAR--TULAATDIGGID--GYSLAAAFWTMGFAIYVGEIGPC VAP-RPVVLAYLLLPLAALAR----FAAATWPETYVAAMLSAAALWVLAFALFALSSAPA : : : : :: * : : : : ::: : *

NmNnrs NGNNRS PsORF396 PsNnrS RcNnrS RcNnrS

LIRPRSDGRP--G LIRPRSDGRP--G LUAARUDGHP--G LCRPRUDGHP--G LLRPSLGRTPSKG FLGPR--QTP--: *

Fig. 5.

784

J.P. Shapleigh

expression of the nar genes. Because nitrate respiration leads to the production of the other denitrification intermediates, it can be difficult in wild type cells to demonstrate that nitrate is the effector for only a limited set of genes. Evidence for a nitrate stimulon was then demonstrated by experiments using nitrite reductase-deficient cells. These experiments showed that nitrate alone was not sufficient to activate expression of genes whose products are required for reduction of the other nitrogen oxides (Tosques, 1997; Korner, 1993). Nor does nitrite alone cause significant induction of nitrate reductase. However, there is evidence of cross talk between the nitrate and nitrous oxide stimulons in P. stutzeri and P. denitrificans (Baumann, 1996; Korner, 1989), although it is difficult to rationalize why this occurs. Experiments monitoring gene expression in relation to oxygen concentration show that the nar genes and nitrate reductase are expressed at higher oxygen levels than the nir and nor genes (Baumann, 1996). Expression of nos occurs at similar oxygen levels. The molecular mechanisms required for nar activation have not been extensively studied in a denitrifying bacterium. A pair of two component sensor-regulators is responsible for regulating nar in E. coli. The function of these proteins has been extensively studied and has been reviewed in Darwin (1996). A nitrate sensor (NarL) and its response regulator (NarX) have been characterized in P. stutzeri. However, their deletion did not affect denitrification (Hartig, 1999). This has led to the suggestion that there is a nitrateresponsive system that functions independently of the NarXL system. Nitric Oxide Stimulon Even before isolation of the proteins regulating the genes encoding the nitrite and nitric oxide reductases, it had been observed that the expression of nitric acid reductase in nitrite reductase mutants was negatively affected. This dependence of nitric oxide reductase expression on the activity of nitrite reductase has been demonstrated in many denitrifiers (Ye, 1992; Tosques, 1997; de Boer, 1994; Zumft, 1994). The decrease in nor gene expression in nitrite reductase mutants appears indirect, as any mutation that leads to a loss of nitrite reductase activity has a negative affect on nor expression (de Boer, 1994; Zumft, 1994). The expression of nir genes is also dependent on nitrite reductase activity (Tosques, 1997). The expression of nir is not directly dependent on nitric oxide reductase, but the accumulation of nitric oxide in nitric oxide reductase mutants probably affects nitrite reductase activity (Kwiatkowski, 1996; Zumft, 1994). The observation that it is nitrite reductase activity not nitrite reductase per se that is

CHAPTER 1.23

required for the expression of nir and nor genes suggests that a product of nitrite reduction is required for gene expression. An obvious candidate for the likely effector is nitric oxide, a possibility consistent with the observation that addition of nitric oxide generators to nitrite reductase-deficient strains results in expression of both nir and nor genes (Kwiatkowski, 1996). Moreover, trapping of nitric oxide by hemoglobin decreases expression of nir and nor genes (Kwiatkowski, 1996). The accumulated evidence strongly indicates that it is the production of nitric oxide that stimulates expression of those genes in the nitric oxide stimulon. However, it is also possible that a derivative of nitric oxide may be the actual signal. The role of nitric oxide as a signal molecule in humans is well known (Schmidt, 1994). The first use of nitric oxide as a signal molecule by a living organism, however, was likely by denitrifying bacteria. It is easy to rationalize why nitric oxide or some derivative serves as a signal molecule for denitrifiers. Denitrifying bacteria must keep the steady state levels of nitric oxide low to minimize potential cytotoxic reactions. As nitric oxide acts to regulate expression of the genes whose products are responsible for establishing the steady state levels of nitric oxide, the possibility nitric oxide might accumulate to cytotoxic levels is abated. Also, as it reacts rapidly with oxygen, nitric oxide will accumulate to the levels required to activate gene expression only when oxygen tension is low. This permits a single molecule to be used as an indicator of both oxygen and nitrogen oxide concentrations in the environment. The direct control of nitric oxide levels by oxygen may explain why nitrite reductase is expressed at lower oxygen concentrations than nitrate and nitrous oxide reductase. In several denitrifiers, a gene has been isolated whose product directly regulates nir and nor expression but not nar or nos genes. The regulation of nir and nor by a single regulatory protein not involved in regulating expression of the other nitrogen oxide reductases is consistent with studies of gene expression indicating three separate stimulons. Not surprisingly, this gene has been given a different name in each bacterium. In P. denitrificans, it is designated nnr (van Spanning, 1995), nnrR in R. sphaeroides (Tosques, 1996), dnr in P. aeruginosa (Arai, 1995) and dnrD in P. stutzeri (Hartig, 1999). The family of proteins encoded by these genes will be referred to as the Nnr family in this review. Recent sequencing efforts have also identified likely orthologs in Synechocystis strain PCC 6803 (Kaneko, 1996), Pseudomonas sp. G-179 (Bedzyk, 1999) and R. capsulatus (Rhodobacter, 1999). All of these proteins are members of the

CHAPTER 1.23

The Denitrifying Prokaryotes

Fnr/CRP family of transcriptional regulators (Spiro, 1994). Comparison of the sequences of these various proteins reveals little about how they might interact with an effector. Significantly, there are no obvious metal binding motifs in any of these proteins. Fnr and CooA, which are also members of the Fnr/CRP family, have metal centers that are targets for effector interaction. Fnr from E. coli binds an iron-sulfur center that apparently undergoes structural changes as the oxygen concentration in the cell changes (Popescu, 1998). CooA from Rhodospirillum rubrum binds a heme protein that acts as a carbon monoxide sensor (Shelver, 1997). Current data is not consistent with members of the Nnr family containing any type of metal center. Phylogenetic analysis of the Nnr family reveals that the relatedness of the proteins does not coincide with relatedness predicted by 16S rRNA analysis. For example, R. sphaeroides is closely related to both R. capsulatus and P. denitrificans. The Nnr from P. denitrificans and the putative Nnr ortholog identified in R. capsulatus have significant identity but have only limited similarity to NnrR from R. sphaeroides (Fig. 6). This suggests species relatedness is not the major factor controlling the degree of relatedness of the members of the Nnr family. Based on available data, a model can be presented in which nitrite produced by nitrate reductase activity is reduced by nitrite reductase, and some compound (most likely nitric oxide or a nitric oxide derivative) activates the transcriptional regulator resulting in expression of genes in the nitric oxide stimulon. Even though this model may be generally correct, differences appear in the regulation of the expression of various members of the Nnr family. In R. sphaeroides, NnrR appears to be constitutively expressed but may be negatively autoregulated (Tosques, 1996). There is no evidence of negative autoregulation of Nnr in P. denitrificans (van Spanning, 1995). Expression of Dnr in P. may be regulated

ScNnr PaDnr PdNnr

RcNnr G179Nnr

243NnrR 241 NnrR Fig. 6.

785

by another member of the Fnr/Crp family, Anr, an apparent ortholog of Fnr in E. coli (Arai, 1995). This suggests a regulatory hierarchy where under low oxygen tension Anr activates expression of dnr, whose product can then activate expression of nir and nor genes under appropriate conditions. This type of regulatory hierarchy is not present in P. stutzeri (Hartig, 1999). In P. aeruginosa, Dnr and Anr both regulate hemN, which encodes an oxygen-independent coproporphyrinogen oxidase required for heme synthesis (Rompf, 1998). The hemN gene is also coregulated by NnrR and FnrL in R. sphaeroides (Shapleigh, unpublished). Both FnrL and NnrR regulate the gene encoding pseudoazurin in R. sphaeroides (Jain and Shapleigh, unpublished). This dual regulation of selective genes by both a global regulator such as Anr/Fnr and a regulator of a limited set of genes such as Dnr/NnrR suggests that transcriptional activation by the global regulator alone is not sufficient for optimal growth under denitrification conditions. Therefore, denitrifiers have developed mechanisms to ensure sufficient levels of expression of genes whose products are in heavy demand during denitrification and other modes of anaerobic growth. Further, in R. sphaeroides nitrite accumulation negatively affects expression of FnrL regulated genes (Shapleigh, unpublished). This may provide another explanation as to why FnrL and NnrR are dual regulators of specific genes. It seems likely such dual regulation will be observed in other denitrifiers and will probably encompass a larger set of gene targets. Nitrous Oxide Stimulon Evidence for a nitrous oxide stimulon was initially provided by the observation that growth on nitrous oxide strongly stimulates nitrous oxide reductase expression, modestly stimulates expression of the nitrate reductase, and does not stimulate expression of nitrite or nitric oxide reductase (Korner, 1989). The observation that inactivation of the regulator of genes in the nitric oxide stimulon did not affect growth at the expense of nitrous oxide provided additional support for existence of a set of genes whose transcription depends solely on the presence of nitrous oxide. The nature of the genes responsible for regulating nitrous oxide expression is not well defined. One gene suggested to play a role in regulation of nos genes is nosR. Inactivation of nosR in P. stutzeri inhibited expression of nosZ, the nitrous-oxide-reductase structural gene (Cuypers, 1992). NosR is a putative membrane protein containing a cytoplasmic C-terminal domain with two motifs that resemble [Fe-S] containing motifs. This unusual combination of a membrane bound, [Fe-S] protein involved

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in expression of a copper-containing protein increases interest in this protein. It seems unlikely NosR is involved in copper metabolism, as inactivation of nosR results in the inhibition of nosZ transcription (Cuypers, 1992). In contrast, inactivation of nosFDY, suggested to be involved in copper metabolism, does not cause inhibition of nosZ transcription (Zumft, 1990). Other Regulatory Proteins Searches for regulatory proteins required for expression of denitrification genes are only just beginning to identify possible regulators and to determine their physiological roles. Although the majority of the genes discussed in the preceding sections have been found in most denitrifiers, other putative regulatory proteins have been identified in only a single bacterium. One example is nirI, a gene implicated in regulation of expression of nirS, the structural gene for nitrite reductase in P. denitrificans (Genbank accession number AJ001308). Interestingly, the nirI product is similar to NosR, which is required for expression of nos genes. The involvement of NirI in nitrite reductase expression makes it less likely that NosR is involved in nitrous oxide reductase assembly. Instead, NosR and NirI are likely members of a family of proteins involved in regulation of nitrogen oxide metabolism in denitrifiers. Other regulatory proteins have been found clustered with denitrification genes, but their role in denitrification has not been defined. Clearly, our understanding of the regulation of denitrification genes is very limited. The list of the various proteins involved in regulation will expand, no doubt, as additional denitrifiers are characterized in greater detail.

Metabolism of Related Nitrogen Oxides Because of its reactivity, nitric oxide will react with many compounds generating a wide range of different nitrogen oxide containing molecules. Some of these derivatives are more toxic than nitric oxide. For example, nitric oxide can react with the thiol of glutathione to generate Snitrosoglutathione (GSNO), which has been shown to be more toxic for Salmonella typhimurium than nitric oxide (De Groote, 1995). Another very toxic nitric oxide derivative, peroxynitrite, is generated from the reaction of nitric oxide and superoxide (Squadrito, 1998). There has been a great deal of interest in the interaction of these types of compounds with pathogenic bacteria inasmuch as they are generated as part of the host defense mechanism during infection. These compounds could possibly

CHAPTER 1.23

be generated during denitrification as well. If so, denitrifiers have probably developed mechanisms to mitigate the potentially cytotoxic effects of such derivatives. Despite the paucity of work on the metabolism of nitric oxide derivatives by denitrifiers, some data suggest that denitrifiers are useful models for understanding how cells mitigate nitric oxide toxicity. In experiments assessing the sensitivity of R. sphaeroides strains to GSNO, those strains with nitrite reductase activity were completely resistant to levels of GSNO to which S. typhimurium exhibited sensitivity (Wu, 1998). Comparable assays on a naturally occurring nitrite reductase-deficient strain of R. sphaeroides demonstrated that its sensitivity to GSNO was similar to that exhibited by S. typhimurium. There was no indication that the resistant strain had any special capacity to degrade GSNO suggesting the likely modification of a GSNOsensitive target in the resistant strain. The results of this study need to be extended to other denitrifiers to determine whether general resistance to GSNO is intrinsic to denitrifiers. Also, it will be interesting to determine if denitrifying bacteria have any resistance mechanisms to other toxic nitric oxide derivatives such as peroxynitrite. Probing the molecular mechanisms denitrifiers have developed for resistance to nitric oxide derivatives is of broad scientific interest and is further justification for the study of the diverse group of bacteria linked by their shared capacity to reduce nitrate and nitrite to gaseous nitrogen oxides and nitrogen gas. Acknowledgments. I would like to thank W. J. Payne for critically reading the manuscript. I would also like to thank him for nurturing my interest in microbiology and introducing me to denitrification. I would also like to thank my colleagues working in this field for their contributions. The work in my laboratory is funded by the U.S. Department of Energy.

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Bonin, P., and Gilewicz, M. 1991. A direct demonstration of co-respiration of oxygen and nitrogen oxides by Pseudomonas nautica: some spectral and kinetic properties of the respiratory components. FEMS Microbiol. Lett. 80:183–188. Braun, C., and Zumft, W. G. 1992. The structural genes of the nitric oxide reductase complex from Pseudomonas stutzeri are part of a 30-kilobase gene cluster for denitrification. J. Bacteriol. 174:2394–2397. Carter, J. P., Hsiao, Y. H., Spiro, S., and Richardson, D. J. 1995. Soil and sediment bacteria capable of aerobic nitrate respiration. Appl. Environ. Microbiol. 61:2852– 2858. Chan, Y.-K., McCormick, W. A., and Watson, R. J. 1997. A new nos gene downstream from nosDFY is essential for dissimilatory reduction of nitrous oxide by Rhizobium (sinorhizobium) meliloti. Microbiol. 143:2817–2824. Chang, C. K., Timkovich, R., and Wu, W. 1986. Evidence that heme d1 is a 1,3 porphyrindione. Biochemistry 25:8447– 8453. Cheesman, M. R., Zumft, W. G., and Thomson, A. J. 1998. The MCD and EPR of the heme centers of nitric oxide reductase from Pseudomonas stutzeri: evidence that the enzyme is structurally related to the heme-copper oxidase. Biochemistry 37:3994–4000. Coyne, M. S., Arunakumari, A., Averill, B. A., and Tiedje, J. M. 1989. Immunological identification and distribution of dissimilatory heme cd1 and nonheme copper nitrite reductases in denitrifying bacteria. Appl. Environ. Microbiol. 55:2924–2931. Cramm, R., Siddiqui, R. A., and Friedrich, B. 1997. Two isofunctional nitric oxide reductases in Alcaligenes eutrophus H16. J. Bacteriol. 179:6769–6777. Cutruzzola, F., Arese, M., Grasso, S., Bellelli, A., and Brunori, M. 1997. Mutagenesis of nitrite reductase from Pseudomonas aeruginosa: tyrosine-10 in the c heme domain is not involved in catalysis. FEBS Lett. 412:365– 369. Cuypers, H., Viebrock-Sambale, A., and Zumft, W. G. 1992. NosR, a membrane-bound regulatory component necessary for expression of nitrous oxide reductase in denitrifying Pseudomonas stutzeri. J. Bacteriol. 174: 5332–5339. Darwin, A. J., and Stewart, V. J. 1996. The NAR modulon systems:nitrate and nitrite regulation of anaerobic gene expression. Lin, E. C. C. and Lynch, A. S. Regulation of gene expression in Escherichia coli. R. G. Landes Co.. Austin, TX. de Boer, A. P., Reijnders, W. N., Kuenen, J. G., Stouthamer, A. H., and van Spanning, R. J. 1994. Isolation, sequencing and mutational analysis of a gene cluster involved in nitrite reduction in Paracoccus denitrificans. Antonie Van Leeuwenhoek. 66:111–127. De Boer, A. P. N., Reijnders, W. N. M., Van der Oost, J., Stouthamer, A. H., and Van Spanning, R. J. M. 1996. Mutational analysis of the nor gene cluster encoding nitric oxde reductase from Paracoccus denitrificans. Eur. J. Biochem. 242:592–600. De Groote, M. A., Granger, D., Xu, Y., Campbell, G., Prince, R., and Fang, F. C. 1995. Genetic and redox determinants of nitric oxide cytotoxicity in a Salmonella typhimurium model. Proc. Natl. Acad. USA 92:6399–6403. Denariaz, G., Payne, W. J., and Legall, J. 1991. The denitrifying nitrite reductase of Bacillus halodenitrificans. Biochim. Biophys. Acta 1056:225–232.

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Sweerts, J. P. R. A., DeBeer, D., Nielsen, P., Verduow, H., Heuvel, J. C. V., Cohen, Y., and Cappenberg, T. E. 1990. Denitrification by sulfur oxidizing Beggiatoa spp. mats on freshwater sediments. Nature 344:762– 763. Teraguchi, S., and Hollocher, T. C. 1989. Purification and some characteristics of a cytochrome c-containing nitrous oxide reductase from Wolinella succinogenes. J. Biol. Chem. 264:1972–1979. Timmer-ten Hoor, A. 1975. A new type of thiosulphateoxidizing, nitrate reducing microorganism: Thiomicrospira denitrificans sp.nov. Neth. J. Sea Res. 9:344– 350. Tosques, I. E., Kwiatkowski, A. V., Shi, J., and Shapleigh, J. P. 1997. Characterization and regulation of the gene encoding nitrite reductase in Rhodobacter sphaeroides 2.4.3. J. Bacteriol. 179:1090–1095. Tosques, I. E., Shi, J., and Shapleigh, J. P. 1996. Cloning and characterization of nnrR, whose product is required for the expression of proteins involved in nitric oxide metabolism in Rhodobacter sphaeroides 2.4.3. J. Bacteriol. 178:4958–4964. Tsukihara, T., Aoyama, H., Yamashita, E., Tomizaki, T., Yamaguchi, H., Shinzawa-Itoh, K., Nakashima, R., Yaono, R., and Yoshikawa, S. 1995. Structures of metal sites of oxidized bovine heart cytochrome c oxidase at 2.8 Å. Science 269:1069–1074. Usuda, K., Toritsuka, N., Matsuo, Y., Kim, D. H., and Shoun, H. 1995. Denitrification by the fungus Cylindrocarpon tonkinense: anaerobic cell growth and two isozyme forms of cytochrome P-450nor. Appl. Env. Microbiol. 61:883–889. van der Oost, J., de Boer, A. P. N., De Gier, J.-W. L., Zumft, W. G., Stouthamer, A. H., and van Spanning, R. J. M. 1994. The heme-copper oxidase family consists of three distinct types of oxidases and is related to nitric oxide reductase. FEMS Microbiol. Lett. 121:1–9. van Niel, E. W., Braber, K. J., Robertson, L. A., and Kuenen, J. G. 1992. Heterotrophic nitrification and aerobic denitrification in Alcaligenes faecalis strain TU. Antonie von Leeuwenhoek. 62:231–237. van Schie, P. M., and Young, L. Y. 1998. Isolation and characterization of phenol-degrading denitrifying bacteria. Appl. Env. Microbiol. 64:2432–2438. van Spanning, R. J. M., De Boer, A. P. N., Reijnders, W. N. M., Westerhoff, H. V., Stouthamer, A. H., and Van Der Oost, J. 1997. FnrP and NNR of Paracoccus denitrificans are both members of the FNR family of transcriptional activators but have distinct roles in respiratory adaptation in response to oxygen limitation. Mol. Microbiol. 23:893–907. van Spanning, R. J. M., DeBoer, A. P. N., Reijnders, W. N. M., Spiro, S., Westerhoff, H. V., Stouthamer, A. H., and Van der Oost, J. 1995. Nitrite and nitric oxide reduction in Paracoccus denitrificans is under the control of NnrR, a regulatory protein that belongs to the FNR family of transcriptional activators. FEBS Lett. 360:151–159. Verkhovskaya, M. L., Garcia-Horsman, A., Puustinen, A., Rigaud, J. L., Morgan, J. E., Verkhovsky, M. I., and Wikstrom, M. 1997. Glutamic acid 286 in subunit I of cytochrome bo3 is involved in proton translocation. Proc Natl Acad Sci USA. 94:10128–10131. Viebrock, A., and Zumft, W. G. 1988. Molecular cloning, heterologous expression, and primary structure of the structural gene for the copper enzyme nitrous oxide

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reductase from denitrifying Pseudomonas stutzeri. J. Bacteriol. 170:4658–4668. Volkl, P., Huber, R., Drobner, E., Rachel, R., Burggraf, S., Trincone, A., and Stetter, K. O. 1993. Pyrobaculum aerophilum sp. nov., a novel nitrate-reducing hyperthermophilic archaeum. Appl. Environ. Microbiol. 59:2918– 2926. Vollack, K. U., Xie, J., Hartig, E., Romling, U., and Zumft, W. G. 1998. Localization of denitrification genes on the chromosomal map of Pseudomonas aeruginosa. Microbiol. 144:441–448. Vorholt, J. A., Hafenbradl, D., Stetter, K. O., and Thauer, R. K. 1997. Pathways of autotrophic CO2 fixation and of dissimilatory nitrate reduction to N2O in Ferroglobus placidus. Arch. Microbiol. 167:19–23. Warnecke-Eberz, U., and Friedrich, B. 1993. Three nitrate reductases activities in Alcaligenes eutrophus. Arch. Microbiol. 159:405–409. Wharton, D. C., and Gibson, Q. C. 1976. Cytochrome oxidase from Pseudomonas aeruginosa IV. Reaction with oxygen and carbon monoxide. Biochim. Biophys. Acta 292:611–620. Williams, P. A. Fülöp, V., Garman, E. F., Saunders, N. F., Ferguson, S. J., and Hajdu, J. 1997. Haem-ligand switching during catalysis in crystals of a nitrogen-cycle enzyme. Nature 389:406–412. Wu, Q., Storrier, G. D., Wu, K. R., Shapleigh, J. P., and Abru__, H. D. 1998. Electrocatalytic reduction of Snitrosoglutathione at electrodes modified with an electropolymerized film of a pyrrole derived viologen system and their application to cellular S-nitrosoglutathione determinations. Anal. Biochem. 263:102–112. Yabuuchi, E., Kosako, Y., Oyaizu, H., Yano, I., Hotta, H., Hashimoto, Y., Ezaki, T., and Arakawa, M. 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. 36:1251–1257. Ye, R. W., Arunakumari, A., Averill, B. A., and Tiedje, J. M. 1992. Mutants of Pseudomonas fluorescens deficient in dissimilatory nitrite reduction are also altered in nitric oxide reduction. J. Bacteriol. 174:2560–2564.

CHAPTER 1.23 Ye, R. W., Averill, B. A., and Tiedje, J. M. 1994. Denitrification: production and consumption of nitric oxide. Appl. Env. Microbiol. 60:1053–1058. Yoneyama, H., and T. Nakae, T. 1996. Protein C (OprC) of the outer membrane of Pseudomonas aeruginosa is a copper-regulated channel protein. Microbiol. 142:2137– 2144. Yoshinari, T. 1980. N2O reduction by Vibrio succinogenes. Appl. Env. Microbiol. 39:81–84. Yoshinari, T., and Knowles, R. 1976. Acetylene inhibition of nitrous oxide reduction by denitrifying bacteria. Biochem. Biophys. Res. Comm. 69:705–710. Zeilstra-Ryalls, J. H., and S. Kaplan, S. 1995. Aerobic and anaerobic regulation in Rhodobacter sphaeroides 2.4.1: the role of the fnrL gene. J. Bacteriol. 177:6422– 6431. Zhang, C.-s., Hollocher, T. C., Kolodziej, A. F., and OrmeJohnson, W. H. 1991. 1991. Electron paramagnetic resonance observations on the cytochrome c-containing nitrous oxide reductase from Wolinella succinogenes. J. Biol. Chem. 266:2199–2202. Zumft, W. G. 1997. Cell biology and molecular basis of denitrification. Microbiol. Mol. Biol. Rev. 61:533–616. Zumft, W. G., Braun, C., and Cuypers, H. 1994. Nitric oxide reductase from Pseudomonas stutzeri: Primary structure and gene organization of a novel bacterial cytochrome bc complex. Eur. J. Biochem. 219:481–490. Zumft, W. G. A., Dreusch, A., Lochelt, S., Cuypers, H., Friedrich, B., and Schneider, B. 1992. Derived amino acid sequence of the nosZ gene (respiratory N2O reductase) from Alcaligenes eutrophus, Pseudomonas aeruginosa and Pseudomonas stutzeri reveal potential coppper-binding sites. Eur. J. Biochem. 208:31–40. Zumft, W. G., and Matsubara, T. 1982. A novel kind of multicopper protein as terminal oxidoreductase of nitrous oxide respiration in Pseudomonas perfectomarinus. FEBS Lett. 148:107–112. Zumft, W. G. A., Viebrock-Sambale, A., and Braun, C. 1990. Nitrous oxide reductase from denitrifying Pseudomonas stutzeri: genes for copper-processing and properties of the deduced products, including a new member of the family of ATP/GTP-binding proteins. Eur. J. Biochem. 192:591–599.

Prokaryotes (2006) 2:793–817 DOI: 10.1007/0-387-30742-7_24

CHAPTER 1.24 gn i x i F-negor t i n iD

se toyrakorP

Dinitrogen-Fixing Prokaryotes ESPERANZA MARTINEZ-ROMERO

Introduction Dinitrogen fixation, the biocatalytic conversion of gaseous nitrogen (N2) to ammonium, is an exclusive property of prokaryotes. The enzymes responsible for this reaction are nitrogenases. Proof that bacteria associated with leguminous plants can fix atmospheric N2 (making it available to the plants for growth) was first reported in 1888 (reviewed in Quispel, 1988). Nitrogen fixation is the most important way N2 from air enters biological systems and therefore it is a key step in the nitrogen cycle. From a practical point of view, the importance of the process rests with its ability to reduce the chemical fertilization of crops, even under conditions of environmental stress (Bordeleau and Prévost, 1994; Zahran, 1999). Indeed, agronomically important crops such as soybean, alfalfa, pea, clover and bean obtain substantial amounts of their nitrogen from bacterial N2 fixation. One of the long-term goals of N2 fixation research is to select or engineer major cereal crops such as rice, maize and sugarcane so they can satisfy the bulk of their nitrogen requirements, either indirectly by association with N2-fixing bacteria, or directly by insertion of N2-fixing genes into the plant. Many diazotrophs (di = two, azote = nitrogen; trophs = eaters: dinitrogen fixers) are found to be associated with the roots of plants where they exchange fixed nitrogen for the products of photosynthesis. Plants associated with N2 fixers can grow in very poor soils and swamps (Koponen et al., 2003) and be used successfully for soil remediation. Nowadays industrial fixation of atmospheric N2 exceeds the amount estimated to be produced by biological nitrogen fixation each year (Karl et al., 2002) and increased nitrogen (N) deposition seems to be responsible for loss of biodiversity and plant species extinction (Stevens et al., 2004). Biological N2 fixation is still the main source of nitrogen in soil, marine environments such as oligotrophic oceanic waters (where dissolved fixed-nitrogen content is extremely low; Zehr et al., 1998; Staal et al., 2003), subtropical

and tropical open ocean habitats (Karl et al., 2002), and hydrothermal vent ecosystems (Mehta et al., 2003). N2 fixation in coastal marine environments may diminish because of habitat destruction and eutrophication (Karl et al., 2002). Dinitrogen fixation may be a major nitrogen source for supporting primary and secondary production of biomass in Antarctic freshwater and soil habitats (Olson et al., 1998) and has been reported to occur in moss carpets of boreal forests (DeLuca et al., 2002) and in woody debris (Hicks et al., 2003). Dinitrogen fixation by bacteria inside insect gut helps to compensate termites for their nitrogen-poor diet (Kudo et al., 1998; Nardi et al., 2002). N2-fixing prokaryotes inhabit a wide range of exterior environments (including soils, seas, and the oceans) and interior environments (including insects, cow rumena, human intestines, and feces; Bergersen and Hipsley, 1970), and even printing machines and paper-making chemicals (Vaisanen et al., 1998). Nevertheless, the presence of a N2-fixing bacterium is not evidence for the occurrence of N2 fixation. On the basis of N balance analyses, N2 fixation seemed to account for excess N in humans with a low N diet, and N-fixing bacteria were obtained from their guts (Bergersen and Hipsley, 1970; Oomen and Corden, 1970). Dinitrogen fixers are encountered in Bacteria and in some groups of Archaea. The number of nitrogen-fixing phyla or lineages within the domain Bacteria increased from 5 to 6 when nitrogen-fixing bacteria were discovered within the Spirochaetes (Lilburn et al., 2001). The inventory of the phyla containing nitrogen-fixing bacteria is probably still far from complete but enlarging, as with the report of a strain of Verrucomicrobium that is reported to have nitrogen fixation genes (Rodrigues et al., 2004). Lists of N2-fixing prokaryotes have been published (Young, 1992; Phillips and Martínez-Romero, 2000), and new nitrogen-fixing species are continuously being described (Chen et al., 2001; Moulin et al., 2001; Distel et al., 2002; Von der Weid et al., 2002; Bianciotto et al., 2003; Rosenblueth et al., 2004). Nevertheless knowledge of

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N2 fixers is limited, and some not yet identified N2 fixers could be found among the novel bacterial divisions that are mostly unculturable (Rappéand Giovannoni, 2003). The distribution of N2 fixers among the prokaryotes is patchy (Young, 1992). They constitute restricted groups within larger bacterial clusters. The existence of non-fixers that are closely related to fixers has been explained by the loss of N2 fixation genes or by the lateral transfer of these genes among bacterial lineages (Normand and Bousquet, 1989; Vermeiren et al., 1999). Nitrogen fixation is an energy costly process, which may explain why nitrogen fixation was lost in many bacterial lineages when not needed. The possession of N2fixing genes does not confer a selective advantage to bacteria in nitrogen-rich environments, as is the case where fixed nitrogen is added to agricultural fields. Application of ammonium sulfate reduced the number of Azotobacter in the plant rhizosphere, and when compared with plants fertilized with both nitrogen and phosphorus, maize treated with phosphate alone had increased nitrogenase activity (Dö bereiner, 1974). Similarly, very few or no Gluconacetobacter diazotrophicus microorganisms were isolated from sugarcane plants from heavily fertilized areas (Fuentes-Ramírez et al., 1993; Muthukumarasamy et al., 1999), and, perhaps as a result of chemical nitrogen fertilization, the bacterial population had very limited genetic diversity (Caballero-Mellado and Martínez-Romero, 1994; Caballero-Mellado et al., 1995). Subsequently, sugarcane colonization by A. diazotrophicus was found to be inhibited in plants supplied with chemical nitrogen fertilizer (Fuentes-Ramírez et al., 1999). Another effect of adding fixed nitrogen (diminished genetic diversity of Rhizobium from Phaseolus vulgaris bean nodules) was observed when the plants were treated with the recommended level of chemical nitrogen (Caballero-Mellado and MartínezRomero, 1999). The complete genome sequence of the Archaeon Methylobacterium thermoautotrophicum was reported in 1997 revealing the presence of nif genes (Smith et al., 1997), but N2 fixation could not be demonstrated in this strain (Leigh et al., 2000). The sequences of the genomes of the legume-nodulating bacteria belonging to the genera of Mesorhizobium (Kaneko et al., 2000), Sinorhizobium (Galibert et al., 2001) and Bradyrhizobium (Kaneko et al., 2002) revealed contrasting chromosome sizes and highly diverging genomes. A common ancestor of Mesorhizobium and Sinorhizobium was deduced to exist nearly 400 million years ago (Morton, 2002). One of the most novel areas in nitrogen fixation research is genomics, and for sure many N2-fixing bacteria will be used for the determination of

CHAPTER 1.24

their whole genome sequence in the near future. Post-genomic studies are already on course as well.

Diazotroph Isolation and Conditions for N2 Fixation N2-fixing bacteria are normally isolated in N-free media. Whether a microorganism is a N2 fixer is not easy to determine. In the past, claims for many fixers were shown to be erroneous, mainly because fixers were recognized by their ability to grow in nitrogen-free media. However, traces of fixed nitrogen in the media sometimes accounted for the bacterial growth. At other times, oligotrophic bacteria and fungi, which can grow on nitrogen-free media, have been incorrectly reported to be N2-fixing organisms. These microorganisms appear to meet their nitrogen requirements by scavenging atmospheric ammonia (Postgate, 1988). Photosynthetic bacteria have been known for more than 100 years, but the capacity of some of these bacteria to fix N2 was not recognized until much later. Microorganisms may fix N2 under special conditions that may not be readily provided in the laboratory. For example, nitrogenases are inactivated in the presence of oxygen, and different levels of oxygen seem to be optimal for different N2-fixing organisms. Also, some bacteria (e.g., some Clostridium) fix N2 only in the absence of oxygen. In other cases, fixation may require specific nutritional conditions or a differentiation process or both. A remarkable case is the differentiation process of Rhizobium to form N2-fixing bacteroids (Bergersen, 1974; Glazebrook et al., 1993) inside plant root or stem nodules. Bradyrhizobium species can fix N2 both in plant nodules and in vitro, when provided with succinic acid and a small amount of fixed nitrogen (Phillips, 1974). To fix N2, bacteria belonging to the genus Azoarcus (obtained from Kallar grass and more recently also from rice plants) require proline, undergo differentiation, and form a structure called a “diazosome” (Karg and Reinhold-Hurek, 1996). Stimulated by plants, cyanobacteria differentiate into N2-fixing heterocysts that protect nitrogenase from oxygen (Wolk, 1996). Light was found to induce circadian rhythms of N2 fixation in the cyanobacterium Synechococcus (Chen et al., 1993). Wheat germ agglutinins were found to stimulate N2 fixation by Azospirillum, and a putative receptor of this agglutinin was found in the Azospirillum capsule. The stimulus generated from the agglutinin-receptor interaction led to elevated transcription of both structural and regulatory nitrogen-fixation genes (Karpati et al., 1999).

CHAPTER 1.24

Methods for Detecting Nitrogen Fixation The methods used to measure biological N2 fixation include the quantification of the total nitrogen difference from Kjeldahl analysis, acetylene reduction, and 15N incorporation or dilution. The acetylene reduction assay has been used for over 30 years to measure nitrogenase activity and as an indicator of N2 fixation (Hardy et al., 1968). These methods have been used both in the laboratory and the field, and improvements of the methods especially for field evaluations have been proposed, including double labeling using 34 S as a control reference (Awonaike et al., 1993). The 15N-based techniques have been thoroughly reviewed (Bergersen, 1980; Hardarson and Danso, 1993). Nitrogenases may reduce other substrates in addition to N2 and this has been the basis for the acetylene reduction assay, which measures N2 fixation activity indirectly. However, the nitrogenase described by Ribbe et al. (1997) does not have the ability to reduce acetylene. In Paenibacillus, N2 fixation has been demonstrated in some cases by the increase in nitrogen measured by the microKjeldahl method but not by acetylene reduction (Achouak et al., 1999). To circumvent the problems of estimating N2 fixation under laboratory conditions, a strategy to detect nitrogenase genes has been successfully followed. This strategy was made possible by identification of conserved signatures (useful as primers for the synthesis of the nitrogenase reductase gene by means of polymerase chain reaction [PCR] amplification) in the structural nif gene sequences, namely nifH, found in many microorganisms (Dean and Jacobson, 1992; Ueda et al., 1995). In other cases, homologous or heterologous probes have been used in hybridization experiments to detect N2 fixers. With some nifH primers containing conserved sequences, alternative nitrogenases may also be amplified but not the nitrogenase (superoxide) that is structurally unrelated to the classical nitrogenase (Ribbe et al., 1997). Thus a search for N2-fixing organisms using a procedure based only on the classical nifH gene would be incomplete. Nevertheless, with nitrogenase DNA primers and PCR synthesis, novel N2-fixing genes may be found. Eight nifH gene types corresponding mainly to those of diazotrophic Proteobacteria were detected in rice root from endophytic or rhizoplane-borne bacteria (Ueda et al., 1995). Remarkably, none of the sequences amplified corresponded to previously described nifH sequences. The nucleotide sequence of one of the types was found to resemble those of the Azoarcus nif genes. Some bacteria in the gut of termites

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also have nifH sequences similar to those obtained from rice roots (Ohkuma et al., 1999). nif genes were found in human and bovine treponemas (Lilburn et al., 2001) but not in the completely sequenced genomes of the spirochetes Treponema pallidum or Borrelia burgdorferi. Few N2-fixing organisms from the oceanic environment have been cultivated and it is estimated that less than 10% of marine diazotrophs are cultivable. Nevertheless, on the basis of the amplification of nitrogenase nifH genes, new N2-fixing organisms have been detected in oligotrophic oceans. Nitrogenase genes characteristic of cyanobacteria and of Alpha- and Betaproteobacteria were obtained, whereas nifH sequences from samples associated with planktonic crustaceans were found to be clustered with the corresponding sequences from either sulfate reducers or clostridia (Zehr et al., 1998). Since knowledge of the nitrogenase gene diversity has improved (over 1500 sequences were available at the time this manuscript was being written), different sets of primers have been designed (Bü rgmann et al., 2004) to better amplify nifH genes directly from DNA extracted from various samples including environmental samples. More diverse diazotrophic populations have been revealed with this approach than with classical microbiological techniques that require culturing of the bacteria (Zehr et al., 1998; Bü rgmann et al., 2004). A different method of N2-fixation detection involves the growth of indicator non-N2-fixing organisms in a co-culture with putative N2-fixing bacteria. Such an approach has the additional advantage of identifying bacteria that not only fix N2 but also can release fixed nitrogen into the environment and thereby have potential use in agriculture. Gluconacetobacter diazotrophicus (Yamada et al., 1997), a N2-fixing isolate from sugarcane, was cultured with the yeast Lipomyces kononenkoae on nitrogen-free medium, and yeast growth was shown to be proportional to the amount of N2 fixed (Cojho et al., 1993).

Distribution of Dinitrogen-Fixing Ability among Prokaryotes Archaea and Bacteria nitrogenases are phylogenetically related (Leigh, 2000), and supposedly the last common ancestor was a N2-fixing organism (Fani et al., 1999). Alternatively, nitrogen fixation could have evolved in methanogenic archaea and subsequently transferred into the bacterial domain (Raymond et al., 2004). Nowadays, only 6 out of 53 currently identifiable major lineages or phyla within the domain Bacteria

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CHAPTER 1.24

have nitrogen-fixing members, namely: Proteobacteria, cyanobacteria, Chlorobi (green nonsulfur), spirochetes and the Gram-positives (Firmicutes and Actinobacteria; Fig. 1). Dinitrogen-fixing organisms have an advantage over non-fixers in N2-deficient but not in N2sufficient environments where the N2 fixers are readily outcompeted by the bulk of microorganisms. The nif genes may be expected to disappear from bacteria that become permanent inhabitants of environments with available fixed N2; this may explain why some non-N2 fixers emerged and are closely related to N2 fixers in phylogenetic trees of bacteria. Even within species of N2 fixers, some strains do not fix N2 perhaps because of the loss of this unique capacity, as is evident in Azotobacter, Beijerinckia (Ruinen, 1974) and Frankia (Normand et al., 1996). In Rhizobium, nif genes and genes for nodule formation may be easily lost concomitantly with the symbiotic plasmid (Segovia et al., 1991). Similarly, nonsymbi-

otic Mesorhizobium strains are found in nature that lack a symbiotic island (Sullivan et al., 1996). N2-fixing species seem to be dominant in Rhodospirillaceae (Madigan et al., 1984), and within the methanogens (in Archaea), nitrogen fixation is widespread (Leigh, 2000). While all Klebsiella variicola isolates were N2-fixing bacteria (Rosenblueth et al., 2004), only 10% of its closest relatives (K. pneumoniae from clinical specimens) had this capacity (Martínez et al., 2004). The N2-fixing capability is unevenly distributed throughout prokaryotic taxa, and N2-fixing bacteria are in restricted clusters among species of non-N2-fixing bacteria. Only a subset of cyanobacterial species are able to fix N2. Gluconacetobacter diazotrophicus and a couple of other N2-fixing species are the only diazotrophs in a larger group comprising Acetobacter, Gluconacetobacter and Gluconobacter (Fuentes-Ramírez et al., 2001). Similarly, among aerobic endospore-forming Firmicutes (Gram-positive

1. V   2. vadin BE97 3.  4.OP3 5.  6.WS3 7.BRC1 8.NKB19 9.Firmicutes 10.OP9 11.WS2 12.Cyanobacteria 13.F  14.OP10 15.SC4 16.Actinobacteria 17.NC10 18. 19.Chlorobi 20.Marine Group A 21.Caldithrix 22.Gemmatimonadetes 23.F   24.Proteobacteria 25.  26. 27.SBR1093 28.  29.OP8 30.OS-K 31. 32.Termite Group 1 33.TM6 34. 35.OP5 36.Spirochaetes 37.ABY1 38.BD1-5 Group 39.OP11 40.WS6 41.TM7 42.Guaymas 1 43.WS5 44.SC3 45.Chloroflexi 46.  47.   48.OP1 49.  50.   51.  52.  53.  

Fig. 1. Relatedness of nifH genes from different organisms according to DNA sequence (after Hurek et al., 1997a; Ueda et al., 1995; Young et al., 1992; Zehr et al., 1995). In parentheses, the Proteobacteria subclass.

CHAPTER 1.24

bacteria), N2 fixers are encountered mainly in a discrete group (defined by cluster analysis from 16S rRNA gene sequences) corresponding to Paenibacillus (Achouak et al., 1999). Among the actinomycetes, N2-fixing Frankia, represented by a diversity of phenotypes from different habitats, are grouped by their 16S rRNA gene sequences (Normand et al., 1996). In Archaea, N2-fixing organisms are found in the methanogen group and in the halophile group within the Euryarchaeota but not in the sulfur-dependent Crenarchaeota (Young, 1992). Pseudomonas spp. were considered unable to fix N2, but recently new isolates have been recognized as N2 fixers. Some isolates, closely related to fluorescent pseudomonads, possess in addition to the FeMo nitrogenase an alternative molybdenum-independent nitrogenase (Loveless et al., 1999; Saah and Bishop, 1999). Dinitrogen-fixing Pseudomonas stutzeri, (previously designated Alcaligenes faecalis) (Vermeiren et al., 1999), is widely used as a rice inoculant in China (Qui et al., 1981). Following rice inoculation, P. stutzeri aggressively colonize the roots, and the nifH gene is expressed in these rootassociated bacteria (Vermeiren et al., 1998). Other reports list different N2-fixing Pseudomonas species that have been isolated from sorghum in Germany (Krotzky and Werner, 1987), from Capparis in Spain (Andrade et al., 1997), and from Deschampsia caespitosa in Finland (Haahtela et al., 1983). The sporadic occurrence of nif genes in Pseudomonas may be explained by the acquisition of these genes by lateral transfer (Vermeiren et al., 1999). Pseudomonas stutzeri strains are known to be naturally competent for DNA uptake (Lorenz and Wackernagel, 1990). Other nifH gene sequences obtained from rice-associated bacteria were in the same cluster as the P. stutzeri nifH gene (Ueda et al., 1995; Vermeiren et al., 1999). The phylogenetic relationship of N2-fixing organisms inferred from the comparative analysis of nif and 16S rRNA gene sequences led Hennecke et al. (1985) to propose that the nifH genes may have evolved in the same way as the organisms that harbor them; a similar conclusion was obtained by Young (1992) from the analysis of a larger number of diazotrophs. Ueda et al. (1995) and Zehr et al. (1995), using different reconstruction methods, reported nifH gene phylogenies in general agreement with the phylogenetic relationships derived from 16S rRNA gene sequences, with some exceptions. A more recent comparison of nifH and 16S rRNA phylogenies has been performed with a very short fragment of the nifH gene. An early possible duplication of nifH and paralogous comparisons make interpretations difficult (see Fig. 3 in Zehr et al., 2003). Four major clusters of nifH are recognized

Dinitrogen-Fixing Prokaryotes

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and functional nitrogenases are found in three of them (Zehr et al., 2003). The phylogenies of nifH genes are continuously revised and updated with novel sequences (including environmental ones) and more robust reconstruction methods. nifH genes from Gammaproteobacteria are found in different groups, as well as those from Betaproteobacteria (Bü rgmann et al., 2004). Anomalies in the phylogenetic position of Betaproteobacteria have been reported as well (Hurek et al., 1997; Minerdi et al., 2001).

Ecology of Dinitrogen-Fixing Prokaryotes The communities of dinitrogen-fixing bacteria in natural environments may be studied with approaches such as the amplification by PCR of the nitrogenase reductase gene (nifH) with nifH primers using environmental DNA, with subsequent analyses by cloning and sequencing, by terminal restriction fragment length polymorphism (T-RFLP; Ohkuma et al., 1999; Tan et al., 2003), or by denaturing gradient gel electrophoresis (DGGE; Muyzer et al., 1993). Hybridization to macro- and microarrays may reveal the presence and frequency of different N2-fixing prokaryotes (Jenkins et al., 2004; Steward et al., 2004). The ecology of the symbiotic N2-fixing soil bacteria that are collectively designated rhizobia, has been comprehensively reviewed by Bottomley (1992), and ecogeographic and diversity reviews of these bacteria have been reported (Martínez-Romero and Caballero-Mellado, 1996; Sessitsch et al., 2002). Additional aspects of Rhizobium ecology in soil also have been reviewed (Sadowsky and Graham, 1998). Frankia symbiosis including some ecological aspects has been reviewed by Baker and Mullin (1992) and by Berry (1994). New molecular approaches have recently enhanced our perception of microorganisms in their natural habitats. By using PCR primers targeted to nitrogenase genes, the description and natural histories of communities of N2-fixing microorganisms may be established more accurately than with traditional microbiological techniques. The fluctuations of marine diazotroph populations have been analyzed with these approaches. The bulk of N2 fixation appears to shift from cyanobacterial diazotrophs in summer to bacterial diazotrophs in fall and winter (Zehr et al., 1995). The heterocystous cyanobacteria are not as efficient fixing nitrogen as the nonheterocystous cyanobacteria at the high temperatures of the tropical oceans (Staal, 2003). The diversity of marine N2 fixers in benthic marine mats was determined from the sequences of nifH genes. The nifH

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sequences obtained were most closely related to those of anaerobes, with a few related to Gammaproteobacteria including Klebsiella and Azotobacter species (Zehr et al., 1995). The role of N2 fixation was examined in microbial aggregates embedded in arid, nutrientlimited and permanent ice covers of a lake area in the Antarctic, and also in mats in soils adjacent to the ice border. Molecular characterization by PCR amplification of nifH fragments and nitrogenase activity measured by acetylene reduction showed a diverse and active diazotrophic community in all the sites of this environment. Nitrogenase activity was extremely low, compared to temperate and tropical systems. Diazotrophs may be involved in beneficial consortial relationships that may have advantages in this environment (Olson et al., 1998). Nitrogen fixation, observed in moderately decayed wood debris, was shown to be stimulated by warm temperatures (Hicks et al., 2003). The diversity of the N2-fixing microorganisms within the symbiotic community in the gut of various termites was studied without culturing the symbiotic microorganisms. Both small subunit (ss) rRNA (Kudo et al., 1998) and nifH genes (Ohkuma et al., 1999) were amplified in DNA extracted from the mixed microbial population of the termite gut. The analysis of the nif clones from diverse termites revealed different sequences in most of the individual termite species. Whereas the nif groups were similar within each termite family, they differed between termite families. Microorganisms from termites with high levels of N2-fixation activity could be assigned to either the anaerobic nif group (clostridia and sulfur reducers) or to the alternative nif methanogen group. Highly divergent nif gene sequences (perhaps not even related to nitrogen fixation) were found in termites that showed low levels of acetylene reduction (Ohkuma et al., 1999). Expression of the N2 fixation gene nifH was evaluated directly by amplifying nifH cDNA from mRNA by reverse transcription (RT)-PCR (Noda et al., 1999). Only the alternative nitrogenase (from anf gene) was preferentially transcribed in the gut of the termite Neotermes koshunensis. The levels of expression of the anf gene were related to the N2 fixation activity recorded for the termites. The addition of Mo (molybdenum) to the termite diet did not repress the expression of the anf genes; however, Mo repression of other anf genes has been described (Noda et al., 1999). Estimates are that the contribution of insectborne nitrogen-fixing bacteria in insects may be up to 30 kg of N/hectare (ha)/year (Nardi et al., 2002). Endosymbionts from marine bivalve species, located in the shipworm gills, are cellulolytic and

CHAPTER 1.24

N2-fixing. They provide cellulolytic enzymes to the host. They are a unique clade in the Gammaproteobacteria related to Pseudomonas and were designated as a new genus and species Teredinibacter turnerae, which fixes nitrogen in microaerobic in vitro conditions (Distel et al., 2002). The arbuscular mycorrhizal fungus (Gigaspora margarita) has been shown to harbor a viable and homogeneous population of endosymbiotic bacteria that has been designated as “Candidatus Glomeribacter gigasporarum” (Bianciotto et al., 1996) related to Betaproteobacteria such as Ralstonia (Bianciotto et al., 2003). In the genomic library of total DNA from the fungal spores, clones carrying the bacterial genes nifD and nifK were identified. Both of these genes were arranged in a similar manner to the corresponding genes in archaea or bacteria and were similar to nitrogenases from different diazotrophs (Minerdi et al., 2001; Minerdi et al., 2002). mRNAs for the nif genes were detected, but whether these endosymbionts fix nitrogen is unknown. Dinitrogen-fixing cyanobacteria form symbioses with diverse hosts such as fungi, bryophytes, cycads, mosses, ferns, and an angiosperm, Gunnera (Bergman et al., 1992). The genome of the cyanobacteria Nostoc (which is a symbiont of cycads, Gunnera and others) may be the largest among those from Prokaryotes, with nearly 10 Mb (Meeks et al., 2001). New symbionts capable of forming nodules in the leguminous plant Lotus corniculatus were obtained in agricultural fields after the lateral transfer of genetic material to native nonsymbiotic soil mesorhizobia (Sullivan et al., 1995; Sullivan et al., 1996). Nonsymbiotic soil rhizobia, which outnumber symbiotic bacteria in some cases (Segovia et al., 1991; Laguerre et al., 1993), have been considered to be potential recipients of symbiotic plasmids. Molecular analyses (including the sequence of DNA fragments of 16S rRNA genes, the fingerprints of digested genomic DNA, and the hybridization patterns to cloned fragments) clearly demonstrated that a large segment of genetic material was acquired by soil Mesorhizobium bacteria (Sullivan et al., 1995) and that the original Mesorhizobium loti strain applied to the soil as an inoculant was the donor of these symbiotic genes. The mobilizable 500-kb DNA fragment has been designated a symbiosis island and it encodes genes for symbiotic N2 fixation (fix genes) as well as those for the synthesis of vitamins (Sullivan et al., 2002). The symbiotic island was integrated into the phenylalanine-tRNA gene (Sullivan and Ronson, 1998). Interestingly, pathogenicity islands in other bacteria range up to 190 kb in size and most are either found adjacent to or integrated

CHAPTER 1.24

within tRNA genes or flanked by insertion sequences (Cheetham and Katz, 1995; Kovach et al., 1996). In M. loti, the symbiotic genes are chromosomally located as in most Mesorhizobium and Bradyrhizobium sp. A similar symbiotic chromosomal region was identified in M. loti (Kaneko et al., 1999) that was later classified as M. huakuii (Turner et al., 2002). Only a few Mesorhizobium species such as M. amorphae possess symbiotic plasmids (Wang et al., 1999b), which are a common characteristic of Rhizobium and Sinorhizobium species (Martínez et al., 1990). The great chromosomal diversity, mainly based on 16S rRNA sequence (Wang and Martínez-Romero, 2000) and on glutamine synthetase (GSII) genes (Wernegreen and Riley, 1999) encountered in M. loti, may be ascribed to the natural occurrence of genetic transfer of symbiotic genes in Mesorhizobium (Sullivan et al., 1996). The range of nodulating bacteria has enlarged. Nodulating Methylobacterium have been reported from Crotalaria nodules (Sy et al., 2001). Surprisingly, some Betaproteobacteria in the genera Burkholderia (Moulin et al., 2001) and Ralstonia (Chen et al., 2001) are capable of nodulating legumes. These bacteria have been classified as Burkholderia phymatum, B. tuberum (Vandamme et al., 2002), B. caribensis (Chen et al., 2003) and Wautersia taiwanensis (previously designated Ralstonia taiwanensis) (Chen et al., 2001; Vaneechoutte et al., 2004). Like Rhizobium and Sinorhizobium spp., these Betaproteobacteria possess symbiotic plasmids that carry nodulation genes (Chen et al., 2003). The similarity of these nod genes to those of the Alphaproteobacteria suggested that lateral transfer of nod genes occurred, most probably from Alpha- to Betaproteobacteria (Moulin et al., 2001; Chen et al., 2003). Similarly the lateral transfer of nod genes has been implied as a possible explanation for the nodulation capacity in Devosia, and a new species has been identified that carries nodD and nifH genes similar to those of R. tropici (Rivas et al., 2002).

Dinitrogen-Fixing Prokaryotes in Agriculture The first industrial production of Rhizobium inoculants began at the end of the nineteenth century. In the absence of nitrogen fertilization, spectacular increases in plant and seed yield may be obtained by inoculation of legumes where the specific rhizobia for the legumes are absent or scarce (Singleton and Tavares, 1986). Factors affecting nodule occupancy by rhizobia inoculants were reviewed by Vlassak and Vanderleyden (1997). Inoculation of soybean is a common practice in Brazil (Hungria et al., 2000) or in the

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United States where production of soybean inoculants is a top priority for inoculant-producing companies (Paau, 1989), and inoculation of cash crops with nitrogen-fixing inoculants is considered a realistic alternative to the ever increasing use of fertilizers. High quality inoculants (whose characteristics were discussed by Maier and Triplett, 1996) as well as the improvement of management systems, are useful not only for agriculture but also for reforestation of devastated areas. Leguminous trees with their corresponding rhizobia have been recommended for many and diverse uses including reforestation, soil restoration, lumber production, cattle forage, and for human food. The so-called “actinorhizal plants” that associate with Frankia are also of great value for reforestation; actinorhizal plants belong to eight families (Baker and Mullin, 1992; Berry, 1994). A high impact goal of nitrogen fixation research has been to extend nitrogen fixation to non-legumes and this has promoted the search for nitrogen fixing bacteria that are associated with agriculturally valuable crops. From a basic research perspective this has increased our knowledge of their diversity. The impact on agriculture and potential as a substitute for the high levels of fertilizer used in intensive agriculture is debatable, and a critical review of the actual contributions of N2 fixation to the amount of fixed N present in cereals and other grasses finds that N2-fixing bacteria in agriculture provide only a limited amount of fixed N. Careful long-term N balance studies would be required to accurately estimate these contributions (Giller and Merckx, 2003). Levels of fixed nitrogen (as low as 5–35 kg N/ha per year) that contribute over the long term to sustain fertility in nonagricultural areas (Stevens et al., 2004) are neglible for present modern intensive agricultural needs but may be of use in traditional, low input small farming systems. Legumes may fix over 200 kg N/ha per year and this is a significant contribution of nitrogen. Conservative values for bacterial fixation in nonlegumes are 20–30 kg N/ha per year, but higher, substantial values have been also estimated (see below). The rate of fixation of the tree Acacia dealbata is considered sufficient to replace the estimated loss due to timber harvesting (May and Attiwill, 2003). Sugarcane and rice are the Gramineae most extensively studied with regard to N2 fixation, but other crops are being studied as well (see below). Sugarcane has been grown for more than 100 years in some areas of Brazil without nitrogen fertilization or with very low nitrogen inputs, and removal of the total harvest has not led to decline in yield and soil nitrogen levels. This observation suggested that N2 fixation may have been the source for a substantial part of the

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nitrogen used by this crop (Dö bereiner, 1961). Alternatively, irrigation water has been implicated as a possible source of N (Giller and Merckx, 2003). From 25–55% (Urquiaga et al., 1989; Yoneyama et al., 1997) or perhaps as much as 60– 80% (Boddey et al., 1991) of the plant N could be derived from associative dinitrogen fixation, but scepticism about the occurrence of high levels of nitrogen fixation has been expressed (Giller and Merckx, 2003). The problems of estimating sugarcane N2 fixation, discussed by Boddey et al. (1995), include different patterns of nitrogen uptake by different sugarcane varieties (Urquiaga et al., 1989), declining 15N enrichment of soil mineral nitrogen, carryovers of nitrogen from one harvest to the next, and differential effects on control plants during the three-year study (Urquiaga et al., 1992). The mean estimates of fixed N2 for two sugarcane hybrids grown in concrete tanks ranged from 170–210 kg N2 fixed/ha (Urquiaga et al., 1992). Correction for micronutrient soil deficiencies and high soil moisture seem to be key conditions that promote N2 fixation in sugarcane plants (Urquiaga et al., 1992). The evidence of large differences in N2 fixation among different sugarcane cultivars is compelling. Dinitrogen-fixing bacteria isolated from the rhizosphere, roots, stems and leaves of sugarcane plants include Beijerinckia, Azospirillum, Azotobacter, Erwinia, Derxia, Enterobacter (reviewed in Boddey et al., 1995), Gluconacetobacter (Cavalcante and Dö bereiner, 1988), and Herbaspirillum (Baldani et al., 1986). Gluconacetobacter diazotrophicus has the capacity to fix N2 at low pH and in the presence of nitrate and oxygen. A G. diazotrophicus nifD mutant that cannot fix N2 has been tested on plants derived from tissue cultures. Plant height was significantly increased by the wildtype strain and not by the mutant strain inoculants, suggesting a positive effect of N2 fixation by G. diazotrophicus on sugarcane (Sevilla et al., 1998). Beneficial effects of G. diazotrophicus inoculation in experimental fields also have been reported (Sevilla et al., 1999), but global N balances were not analyzed. Selected strains of Herbaspirillum were reported to stimulate plant development (Baldani et al., 1999). Gluconacetobacter diazotrophicus (James and Olivares, 1997), Herbaspirillum seropedicae and H. rubrisubalbicans (Olivares et al., 1996) have been clearly shown to colonize sugarcane plants internally. Colonization by G. diazotrophicus was inhibited by nitrogen fertilization (FuentesRamírez et al., 1999). Probably N 2 fixation in sugarcane is performed by a bacterial consortium. Several studies have been carried out on nitrogen balance in lowland rice fields in Thai-

CHAPTER 1.24

land (Firth et al., 1973; Walcott et al., 1977), in Japan (Koyama and App, 1979), and at the experimental fields of the International Rice Research Institute (IRRI) in the Philippines (App et al., 1984; Ventura et al., 1986). These studies report a positive balance with estimates of around 16–60 kg of nitrogen fixed per ha per crop (App et al., 1986; Ladha et al., 1993). In a nitrogen-balance study carried out on 83 wild and cultivated rice cultivars (6 separate experiments, each with 3 consecutive crops), large and significant differences between cultivars were found (App et al., 1986). But other assays showed only a small or nonsignificant contribution of fixed N2 in rice (Watanabe et al., 1987b; Boddey et al., 1995). Many different N2-fixing bacteria have been isolated from rice roots. These include Azotobacter, Beijerinckia (Dö bereiner, 1961), Azospirillum (Baldani and Dö bereiner, 1980; Ladha et al., 1982), Pseudomonas (Qui et al., 1981; Barraquio et al., 1982; Barraquio et al., 1983; Watanabe et al., 1987a; Vermeiren et al., 1999), Klebsiella, Enterobacter (Bally et al., 1983; Ladha et al., 1983), Sphingomonas (described as Flavobacterium in Bally et al., 1983), Agromonas (Ohta and Hattori, 1983), Herbaspirillum spp. (Baldani et al., 1986; Olivares et al., 1996), sulfurreducing bacteria (Durbin and Watanabe, 1980; reviewed in Barraquio et al. [1997] and in Rao et al. [1998]), Azoarcus (Engelhard et al., 1999) and methanogens (Rajagopal et al., 1988; Lobo and Zinder, 1992). The nitrogenase genes of Azoarcus are expressed on rice roots (Egener et al., 1998), and Herbaspirillum seropedicae expresses nif genes in several gramineous plants including rice (Roncato-Maccari et al., 2003). Cyanobacteria have long been used to fertilize agricultural land throughout the world, most notably rice paddies in Asia. Increases in rice plant growth and increases in nitrogen content in the presence of cyanobacteria have been documented by many investigators. Plant promotion may also be related to growth-promoting substances produced by the cyanobacteria (Stewart, 1974). Azolla is a small freshwater fern that grows very rapidly on the surface of lakes and canals. Extensive employment of AzollaAnabaena as a green manure in rice cultivation has been documented. Anabaena, a representative filamentous cyanobacterium, establishes symbioses with a diversity of organisms including Azolla. Unfortunately, various cyanobacteria also produce highly poisonous toxins and some of them are related to the high incidence of human liver cancer in certain parts of China. Highly toxic strains have been found in Anabaena and in other genera of cyanobacteria, and identification of such strains requires sophisticated biochemical tests (Carmichael, 1994).

CHAPTER 1.24

Alternatively, other bacterial species are being tested to promote rice growth, such as the N2fixing Burkholderia vietnamiensis (Gillis et al., 1995). In some agriculture sites in Vietnam, this species has been isolated as the dominant N2fixing bacterium in the rice rhizosphere (Trân Van et al., 1996). Burkholderia vietnamiensis inoculation has resulted in significant increases (up to 20%) in both shoot and root weights in pots and its use in rice inoculation seems highly promising (Trân Van et al., 1994). However, a note of caution has been raised with a proposed moratorium on the agricultural use of B. vietnamiensis, which has a close genetic relationship to human pathogens implicated in lethally infecting patients with cystic fibrosis (Holmes et al., 1998). Detailed molecular analysis may allow for the distinction of pathogenic and environmental isolates (Segonds et al., 1999). For over seven centuries, rice rotation with clover has significantly benefited rice production in Egypt. Clover is normally associated with Rhizobium leguminosarum bv. trifolii that forms N2-fixing nodules in the root of this plant. Surprisingly, strains of this bacterium also were encountered inside the rice plant with around 104–106 rhizobia per gram (fresh weight) of root. These values are within the range of other bona fide endophytic bacteria (Yanni et al., 1997). Promotion of rice shoot and root growth was dependent on the rice cultivar, inoculant strain, and other conditions. Inoculation of rice with a selected strain gives best results in presence of low doses of nitrogen fertilizer. A number of investigators have reported growth stimulation of crops such as wheat and corn inoculated with a R. leguminosarum bv. trifolii strain, but these effects may not be related to N2 fixation (Holflich et al., 1995). In non-legumes (such as Arabidopsis thaliana [a model plant]), penetration of rhizobial strains has been found to be independent of nodulation genes that are normally required for bacterial entry into the legume root (Gough et al., 1996; Gough et al., 1997; Webster et al., 1998; O’Callaghan et al., 1999). This process probably requires cellulases and pectinases (Sabry et al., 1997). Azorhizobium caulinodans, in addition to forming nodules on Sesbania rostrata, has been found to colonize the xylem of its host (O’Callaghan et al., 1999) as well as to colonize wheat (Sabry et al., 1997). In wheat, A. caulinodans promotes increases in dry weight and nitrogen content as compared to uninoculated controls; acetylene reduction activity was also recorded. The interaction between azorhizobia and wheat root resembles the invasion of xylem vessels of sugarcane roots by G. diazotrophicus (James and Olivares, 1997) and Herbaspirillum spp. (Roncato-Maccari et al., 2003) and of

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wheat by Pantoea agglomerans (Ruppel et al., 1992). The xylem vessels may be the site of N2 fixation because they provide the necessary conditions (carbohydrates and low oxygen tension), although the nutrient levels in the xylem have been considered as too low to maintain bacterial growth and N2 fixation (Fuentes-Ramírez et al., 1999; Welbaum et al., 1992). In acreage cultivated using Sesbania rostrata-rice rotation, A. caulinodans survives in the soils and rhizosphere of wetland rice (Ladha et al., 1992). Azorhizobium caulinodans can colonize the rice rhizosphere (specifically around the site of lateral root emergence), penetrate the root at the site of emergence of lateral roots, and colonize subepidermally intercellular spaces and dead host cells of the outer rice root cortex (Reddy et al., 1997). The application of green manure has been an agronomic practice for increasing rice production, and legumes also can be used because of their symbiosis with N2-fixing rhizobia. A large number of species are used both before and after rice culture including Macroptilium atropurpureum, Sesbania and Aeschynomene spp. (Ladha et al., 1992). Owing to their high N2-fixing capacity and their worldwide distribution, flood-tolerant legumes such as Sesbania rostrata have been the focus of research. Sesbania herbacea nodulated by R. huautlense is also a flood-tolerant symbiosis (Wang and MartínezRomero, 2000). Nitrogen fixation in non-legumes is conditioned more by the plant than by the bacteria. Interestingly, aluminum-tolerant plants are more capable of maintaining bacterial nitrogen fixation than plants that are not tolerant (Christiansen-Weniger et al., 1992), maybe because they excrete dicarboxylics that are adequate to support bacterial N2-fixation. N2-fixing bacteria associated to maize include: Azospirillum, Herbaspirillum, Klebsiella (Chelius and Triplett, 2001), Burkholderia vietnamiensis (Trâ n Van et al., 1996), R. etli (GutiérrezZamora and Martínez-Romero, 2001), and the newly described species (Paenibacillus brasilensis; [Von der Weid et al., 2002] and Klebsiella variicola [Rosenblueth et al., 2004]). Klebsiella variicola was also found associated with banana plants (Martínez et al., 2003). Soil type instead of the maize cultivar determined the structure of a Paenibacillus community in the rhizosphere (Araujo de Silva et al., 2003). Sweet potato (Ipomoea batatas) may grow in poor N-soil and associated N-fixation has been considered to contribute N to these plants. By a cultivation-independent approach, bacteria similar to Klebsiella, Rhizobium and Sinorhizobium were inferred to be present as sweet potato endophytes (Reiter et al., 2003).

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Perspectives of Application of Nitrogen Fixation Research The transgenic plants that will herald a revolution in agriculture are those with functional nitrogenase genes that, when expressed, will satisfy all the plant’s nitrogen needs. The source of these genes will be prokaryotic. Research efforts are directed towards the ambitious goal of transforming rice plastids (Potrykus group in Zü rich discussed in Rolfe et al., 1998) and plastids of the alga Chlamydomonas reinhardtii (Dixon et al., 1997; Dixon, 1999). Introduction of additional genes into plants to protect nitrogenase from oxygen damage will be needed. Such approaches could only be based on a profound understanding of N2 fixation biochemistry, gene regulation and organization, as well as the structure and function of nitrogenases. Whether such a goal is feasible is difficult to predict. The identification and selection of plantassociated microorganisms and their genetic improvement is an alternative strategy for obtaining agricultural crops that benefit from prokaryotic N2 fixation. N2 fixation (N2 fixation without nodules) from associated bacteria is being considered as a suitable mode to exploit N2 fixation in non-legumes (Triplett, 1996). Rhizosphere N2 fixation by Rahnella aquatilis has been reported to occur in maize and wheat (Berge et al., 1991), and in other plants (Heulin et al., 1989). Mycorrhiza associate with most plants, and interestingly, bacteria-like organisms with nitrogenase genes have been found to be natural endosymbionts of the mycorrhiza (Minerdi et al., 2002). This association may be exploited to transfer N2 fixation to non-legumes. The genetic improvement of mycorrhiza and bacterial symbionts may constitute a highly efficient system for the provision of fixed nitrogen to the plants. The usefulness of N2-fixing bacteria in bioremediation is also being recognized (Suominen et al., 2000; Prantera et al., 2002). Increased transformation of contaminating polychlorinated biphenyls was obtained with alfalfa inoculated with Sinorhizobium meliloti at 44 days after planting (Mehmannavaz et al., 2002). Dinitrogen fixation may decrease the need for nitrogen required by bacterial consortia used to degrade diesel fuel (Piehler et al., 1999). Novel N2 fixers may be found if the enrichment conditions for their isolation are more varied so as to include aerobic, anaerobic or microaerobic conditions, a variety of carbon sources at varying concentrations (copiotrophic and oligotrophic conditions; Kuznetsov et al., 1979), and media formulations that include or exclude Mo or V. The discovery of a molybdenum-dinitrogenase and a manganese-superoxide

CHAPTER 1.24

oxidoreductase from Streptomyces thermoautothrophicus (Ribbe et al., 1997) opens a new avenue in N2 fixation research. Undoubtedly, other microorganisms containing this nitrogenase have yet to be identified. This nitrogenase may prove to be more amenable for introduction into plants because of its lower energy requirements and its higher tolerance to oxygen. Elevated CO2 levels provided to legumes were found to stimulate N2 fixation indicating that N2 fixation was limited by the availability of photosynthate (Zanetti et al., 1996). Environmental and management constraints to legume growth (basic agronomy, nutrition, water supply, diseases, and pests) are the major limiting factors of N2 fixation in many parts of the world. Crop production on 33% of the world’s arable land is limited by phosphorus availability (Sánchez and Vehara, 1980). Efforts to maximize the input of biologically fixed nitrogen into agriculture will require concurrent approaches, which include the alleviation of phosphorus and water limitation, the enhancement of photosynthate availability, as well as sound agricultural management practices.

Biochemistry and Physiology of Dinitrogen Fixation Although the chemical nature of the primary product of N2 fixation was the subject of debate for many years, the issue was clarified with the use of 15N. All diazotrophs were thought to use the same two-component nitrogenases (consisting of an iron and an molybdenum-iron protein). Alternative nitrogenases were reported subsequently (Hales et al., 1986; Robson et al., 1986) and found in very different bacteria including Anabaena variabilis, Azospirillum brasilense, Clostridium pasteurianum, Heliobacter gestii, Rhodobacter capsulatus, Rhodospirillum rubrum, and bacteria corresponding to Gammaproteobacteria such as Pseudomonas (Saah and Bishop, 1999). Azotobacter vinelandii, an aerobic soil bacterium, was the first diazotroph shown to have three distinct nitrogenases: the classical molybdenum (Mo)-containing nitrogenase (nitrogenase 1), the vanadium (V)containing (nitrogenase 2), and the iron-only nitrogenase (nitrogenase 3; Maynard et al., 1994). The alternative nitrogenases (nitrogenase 2) use V instead of Mo, and this substitution is advantageous under conditions where Mo is limiting (Jacobitz and Bishop, 1992). Similarly, the iron nitrogenase (nitrogenase 3) is expressed only in Mo- and V-deficient, nitrogen-free media. The V-containing nitrogenase produces around three times more hydrogen than the Monitrogenase (Eady, 1996).

CHAPTER 1.24

A Mo-dinitrogenase and a manganesesuperoxide oxidoreductase have been found to couple N2 reduction to the oxidation of superoxide. This nitrogenase is more efficient than the classical enzyme, which requires a fourfold greater input of ATP. This N2-fixing system, which is not sensitive to oxygen, has only been described in Streptomyces thermoautotrophicus (Ribbe et al., 1997), and the genomic DNA of this bacterium does not hybridize to DNA probes for the classical nif genes. Although the overall reactions catalyzed by S. thermoautotrophicus are similar to those of previously characterized nitrogenases (e.g., the production of H2), it is the subunit structure, polypeptides, and inability to reduce acetylene that distinguishes the nitrogenase of this system from other nitrogenases (Ribbe et al., 1997). The currently known dinitrogenase reductases are ca. 63-kDa g2 dimeric iron proteins that contain 4 Fe and 4 S–2 per dimer. In contrast, the St2 protein of S. thermoautotrophicus has been identified as a member of the manganese-superoxide oxidoreductases (SODs) with molecular mass ~48 kDa and no Fe or S–2. Unlike other SODs, St2 cannot convert O– 2 into O2 and H2O2. Some diazotrophs are able to utilize the H2 evolved from N2 fixation via uptake hydrogenases (Evans et al., 1985). These enzymes are found in N2-fixing and non-N2-fixing bacteria and in cyanobacteria. The uptake hydrogenases in Anabaena are present only in heterocysts, which are the specialized N2-fixing cells of cyanobacteria; interestingly, the hydrogenase genes are rearranged during heterocyst differentiation (Carrasco et al., 1995). Hitherto, ammonium has been accepted as the primary product of N2 fixation and as a reactant in the biosynthesis of all nitrogen-containing molecules made by N2-fixing organisms. Because ammonia excretion has been considered a beneficial characteristic enabling N2 fixers to establish symbioses with other organisms such as plants, it has been generally assumed that the ammonium assimilation enzymes are depressed in symbiotic bacteria. However, Bradyrhizobium japonicum, which forms nodules and fixes nitrogen in soybean plants has been shown to excrete alanine preferentially and not ammonium (Waters et al., 1998). Whether this generally occurs in rhizobia is still controversial (Youzhong et al., 2002; Lodwig et al., 2003; Lodwig et al., 2004). The ratio of alanine to ammonia excretion seems to be related to the oxygen concentration and the rate of respiration (Li et al., 1999). For the cyanobacterium Nostoc, which can establish symbiosis with many organisms including Gunnera, ammonia excretion accounts for only 40% of the nitrogen released (Peters and Meeks, 1989). Different plant endophytes have been found to release (excrete) riboflavin during N2 fixation (Phillips et

Dinitrogen-Fixing Prokaryotes

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al., 1999b). Lumichrome, a compound obtained from riboflavin, has been reported to stimulate root respiration and promote alfalfa seedling growth (Phillips et al., 1999a). Production of riboflavin-lumichrome by plant-associated bacteria is favored by a high N-to-C ratio in the media, and possibly N2 fixation also promotes the synthesis of nitrogen-containing compounds (other than ammonia), such as lumichrome, that can benefit plants. Nitrogenase Structure The classical nitrogenase is a complex, two-component metalloprotein composed of an iron (Fe) protein and a molybdenum-iron (MoFe) protein. The properties of nitrogenase have been reviewed (Howard and Rees, 1994; Burgess and Lowe, 1996; Eady, 1996; Seefeldt and Dean, 1997). The iron-molybdenum cofactor (Fe-Moco), the prototype of a small family of cofactors, is a unique prosthetic group that contains Mo, Fe, S, and homocitrate in a ratio of 1 : 7 : 9 : 1, and it is the active site of substrate reduction (Hoover et al., 1989; Kim and Rees, 1992b). All substrate reduction reactions catalyzed by nitrogenase require the sequential association and dissociation of the two nitrogenase components. A great deal of effort to define the structure of nitrogenases has been expended. Azotobacter vinelandii has been suitable for these studies because it produces large amounts of the enzyme, it is amenable to genetic manipulation, and it has nif and nif-associated genes of known sequence (Brigle et al., 1985; Jacobson et al., 1989; Bishop and Premakumar, 1992). A major achievement in the biochemistry of nitrogenases has been the establishment of the structure of the Fe (Georgiadis et al., 1992) and the MoFe proteins (Kim and Rees, 1992b; Bolin et al., 1993; Schindelin et al., 1997) involving high resolution X-ray crystallographic analysis (Peters et al., 1997; Schlessman et al., 1998). A ~2.2 Å resolution has been reported for the Azotobacter vinelandii MoFe-protein (Peters et al., 1997), the A. vinelandii Fe-protein (Av2), and the Clostridium pasteurianum Fe-protein (Schlessman et al., 1998). The knowledge of the Fe protein structure has contributed to understanding how MgATP functions in nitrogenase catalysis. The Fe-protein is a homodimer with two ATP-binding sites, and the nucleotide binding causes conformational changes in the protein. ATP hydrolysis occurs in the transient complex formed between the component proteins. Molecular interactions were proposed from mutagenesis studies of the nitrogenases (Kent et al., 1989; Dean et al., 1990; Scott et al., 1990). Site-specific mutagenesis studies based on the FeMo protein crystal structure (Kim and Rees, 1992a) have been aimed at amino acids related to the FeMo-cofactor (espe-

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CHAPTER 1.24

cially at the residues proposed to be involved in the entry and exit path for substrates, inhibitors and products) and also at those residues involved in FeMo-cofactor insertion during biosynthesis. The spectroscopic and kinetic properties of the resulting mutant proteins are studied (Dilworth et al., 1998). The use of biophysical, biochemical and genetic approaches have facilitated the analysis of the assembly and catalytic mechanisms of nitrogenases. The synthesis of the prosthetic groups of nitrogenases has been a challenge for chemists. The different substrates utilized by the nitrogenases seem to bind to different areas of the FeMo-cofactor (Shen et al., 1997). Nitrogenase structural changes that occur after the formation of the active complex are thought to produce transient cavities within the FeMo protein, which when opened allows the active site to become accessible (Fisher et al., 1998). The FeMo-cofactor also is found associated with the alternative nitrogenase, anf-encoded proteins (AnfDGK; Gollan et al., 1993; Pau et al., 1993). The nifDK genes of Azotobacter vinelandii were fused and then translated into a single large

nitrogenase protein that interestingly has nitrogen fixation activity (Suh et al., 2003). This shows that the MoFe protein is flexible. However a substitution of tungsten for Mo abolished nitrogenase activity (Siemann et al., 2003). Nitrogen Fixation Genes The complete nucleotide sequence of the Klebsiella pneumoniae 24-kb region required for N2 fixation was reported in 1988 (Arnold et al., 1988). Genes for transcriptional regulators were found to cluster contiguously with the structural genes for the nitrogenase components and genes for their assembly. The N2 fixation (nif) genes are organized in seven or eight operons containing the following nif genes: J, H, D, K, T, Y, E, N, X, U, S, V, W, Z, M, F, L, A, B and Q (Fig. 2). The products of at least six N2 fixation (nif) genes are required for the synthesis of the ironmolybdenum cofactor (FeMo-co): nifH, nifB, nifE, nifN, nifQ, and nifV. NifU and NifS might have complementary functions mobilizing the Fe and S respectively needed for nitrogenase metallocluster assembly in A. vinelandii. Notably, some of the gene products required for forma-

Sinorhizobium meliloti nifN

Alphaproteobacteria

H

D

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fixA B C X

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Bradyrhizobium japonicum nifN K

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Klebsiella Gammaproteobacteria nifQ B

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Anabaena vegetative cell nifW X N E

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Cyanobacteria Anabena heterocyst nifW X N E

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Methanococcus maripaludis Archaea

nifH

nifD nifK nifE nifN nifX

nifI1 = gln B1 nifI2 = gln B2 Chlorobium tepidum Green sulfur (Chlorobi)

nifH nifD glnB glnB

nifK

Fig. 2. Arrangements of nif genes in dinitrogen-fixing prokaryotes. The nif gene organization in Methanococcus maripaludis is from Kessler et al. (2001).

CHAPTER 1.24

tion of the Mo-dependent enzyme are also required for maturation of alternative nitrogenases (Kennedy and Dean, 1992). The nifJ gene of Klebsiella is required for N2 fixation, but in the cyanobacterium Anabaena, NifJ is required for N2 fixation only when Fe is limiting (Bauer et al., 1993), whereas in R. rubrum, a NifJ protein does not seem to be required for N2 fixation (Lindblad et al., 1993). The organization of nif genes in Anabaena is unique and different from that of other N2 fixers because nifD is split between two DNA fragments separated by 11 kb. Recombination events are required to rearrange a contiguous nifD gene in N2-fixing cells (Haselkorn and Buikema, 1992; Fig. 2). A detailed analysis of the gene products of nifDK and nifEN (Brigle et al., 1987) revealed a possible evolutionary history involving two successive duplication events. A duplication of an ancestral gene that encoded a primitive enzyme with a low substrate specificity might have occurred before the last common ancestor of all living organisms emerged (Fani et al., 1999). Nitrogenase structural genes are located on plasmids in some bacteria (such as Rahnella aquatilis [Berge et al., 1991], Enterobacter, and Rhizobium spp. [Martínez et al., 1990]) but are chromosomally encoded in the majority of prokaryotes including bradyrhizobia and most mesorhizobia. The repeated sequences clustered around the nif region of the Bradyrhizobium japonicum genome may be involved in recombination thereby facilitating the formation of deletions (Kaluza et al., 1985). In R. etli bv. phaseoli, multiple copies of the nif operon promote major rearrangements in the symbiotic plasmid at high frequency (Romero and Palacios, 1997). Differences in the promoter sequences of the nifH regions in R. etli are correlated with the different levels of nif gene expression (Valderrama et al., 1996). The symbiotic plasmid of R. etli bv. mimosae is closely related to that of bv. phaseoli but its nif gene has a different restriction fragment length polymorphism (RFLP) pattern as revealed by nifH gene hybridization (Wang et al., 1999a). A conserved short nucleotide sequence upstream of genes regulated by oxygen (i.e., an anaerobox) has been detected upstream of Azorhizobium caulinodans nifA (Nees et al., 1988), Bradyrhizobium japonicum hemA, S. meliloti fixL, fixN, fixG, in front of an open reading frame located downstream of S. meliloti fixS, within the coding region of R. leguminosarum bv. viciae fixC, i.e., upstream of the nifA gene and upstream of the fnr gene (fixK-like). Alternative nitrogenase genes, anfH, anfD and anfG (Mo-independent) are found in the termite gut diazotrophs. The sequences of these

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genes are similar to those found in bacteria even though the gene organization with contiguous GlnB-like proteins resembles that found in the Archaea (Noda et al., 1999). The existence of structural genes for three different nitrogenases was revealed when the complete genome sequence of the photosynthetic bacterium Rhodopseudomonas palustris was determined (Larimer et al., 2004). Previously, only Azotobacter sp. was known to possess three nitrogenases. The expression of nif genes of Azotobacter vinelandii was determined directly in soil by PCR amplification of reverse transcribed nifH gene fragments using nifH primers specific for A. vinelandii (Bü rgmann et al., 2003). Regulation of Nitrogen Fixation Genes Since nitrogen fixation is an energy expensive process, it is finely tuned, with transcriptional as well as posttranslational regulation. nif genes are normally not expressed and require transcriptional activation when N is limiting and conditions are appropriate for nitrogenase functioning. If little is known about the extant diazotrophs, less is known about N2 fixation gene regulation from a global phylogenetic perspective. Most studies have been directed to Proteobacteria. For actinobacteria and firmicutes there is almost no information. Cyanobacteria and more recently Archaea were studied and showed very different regulation mechanisms from the ones observed in Proteobacteria. In Archaea, a repressor of nif genes has been identified (Lie and Leigh, 2003) and no nifA has been found in cyanobacteria (Herrero et al., 2001). Novel regulatory elements, their fine interaction, and a huge complexity of regulatory networks are being revealed as the regulation of nitrogen fixation is studied in depth in model bacterial species. The results are revealing a very complicated sequence of regulatory cascades (Dixon, 1998; Nordlund, 2000; Forchhammer, 2003; Zhang et al., 2003). Regulatory elements such as PII (also known as glnB), DRAT (that transfers a ribosyl to nitrogenase and interferes with its activity), and DRAG (that removes the ribosyl) have been found in many diverse nitrogen fixing or non-nitrogen fixing Proteobacteria, Actinobacteria and Archaea (Ludden, 1994; Zhang et al., 2003). Very diverse modes of regulation of nif genes have been described that vary between species or even between strains in a single species (D’hooghe et al., 1995; Girard et al., 2000). Detailed studies have been carried out in Klebsiella pneumoniae, Azotobacter vinelandii, Azospirillum brasilense, Rhodobacter capsulatus, Rhodospirillum rubrum, Sinorhizobium meliloti, Bradyrhizobium japonicum, etc. The most common nitrogenases studied are inactivated by oxygen, and accordingly, the expression

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of nif genes is negatively regulated by high oxygen concentrations. Different oxygen protection mechanisms have been described (reviewed by Vance, 1998). Some of the bacterial diazotrophs share a common mechanism of transcriptional initiation of nif genes using a RNA polymerase holoenzyme containing the alternative sigma factor sN (s54) and the transcriptional activator NifA (Kustu et al., 1989). Regulators of NifA vary among different diazotrophs. Factor sN is competent to bind DNA, but the formation of the open promoter complex (active for transcriptional initiation) is catalyzed by NifA in a reaction requiring nucleoside triphosphate hydrolysis (Lee et al., 1993; Austin et al., 1994). The dual regulation by s54 and NifA may be required to ensure a stringent regulation of nif gene expression, and this may be so because biological N2 fixation represents a major energy drain for the cell. In addition it seems reasonable that nif genes are negatively regulated by ammonia to avoid production of the enzyme in the presence of available fixed nitrogen; accordingly, nitrogenase enzymes are inactivated by ammonia but to a lesser degree in Gluconacetobacter diazotrophicus (Perlova et al., 2003). In vivo DNA protection analysis demonstrated that NifA binds to the upstream activator sequences of nif genes (Morett and Buck, 1988). In the Alpha- and Betaproteobacteria, the activity of NifA is modulated negatively by the antiactivator NifL, which is a flavoprotein. The integrated responses to fixed nitrogen, oxygen, and energy status are mediated via NifL. The oxidized form of NifL inhibits NifA activity. A potential candidate Fe-containing electron donor involved in the signal transduction of NifL may be a flavohemoglobin, which may act as a global intracellular oxygen sensor (Poole et al., 1994). The expression of nifL and nifA in Klebsiella pneumoniae are coupled at the translational level (Govantes et al., 1998). Mutant forms of NifA were obtained that are no longer inhibited by NifL in Azotobacter vinelandii (ReyesRamírez, 2002). In other diazotrophic Proteobacteria, the NifA protein itself senses oxygen probably via a cysteine-rich motif between the central domain and the C-terminal DNA-binding domain (Fischer et al., 1988). Oxygen-tolerant variants of the S. meliloti NifA proteins have been obtained (Krey et al., 1992). Ammonium-insensitive NifA mutants have been reported with modifications involved in the N-terminus of the molecule in Herbaspirillum seropedicae, Azospirillum brasilense and Rhodobacter capsulatus (Souza et al., 1995; Arsene et al., 1996; Kern et al., 1998). In Klebsiella pneumoniae, the nif mRNAs were found to be very stable under conditions

CHAPTER 1.24

favorable to N2 fixation, but the half lives of the nifHDKTY were reduced several fold when adding O2 or fixed nitrogen. A fragment of the nifH sequence is required for the O2-regulation of mRNA stability, and NifY may be involved in the sensing process (Simon et al., 1999). Symbiotic nitrogen fixation shares common elements with free-living nitrogen fixation, but there are substantial differences as well. In Rhizobium, N2 fixation only takes place inside the nodule. Still not well understood is how the plant partner influences the N2-fixing activity of the microsymbiont, and the same is true for termite-diazotroph symbioses as well as for cyanobacteria in plants. In the latter case, the plant seems to stimulate the formation of heterocysts, which are differentiated cells that fix N2 (Wolk, 1996). Even among symbiotic bacteria of legumes (Sinorhizobium, Rhizobium, Azorhizobium and Bradyrhizobium), differences in the fine mechanisms regulating N2 fixation exist and have been reviewed (Fischer, 1994; Kaminski et al., 1998). In S. meliloti, fixLJ (David et al., 1988) gene products belong to a two-component regulatory family of proteins that are responsive to oxygen. FixL is a high affinity oxygen sensor hemoprotein that has kinase-phosphate activity and is involved in phosphorylation of FixJ in microoxic or anoxic conditions (Gilles-Gonzalez et al., 1994). Upon phosphorylation, FixJ binds to the nifA and fixK promoters and allows their transcriptional activation (Waelkens et al., 1992). Nitrogen fixation takes place in heterocysts in some cyanobacteria. Heterocyst differentiation is regulated by HetR, a protease (Haselkorn et al., 1999), and is inhibited by ammonia (Wolk, 1996). The expression of nif genes is also downregulated by ammonium or nitrate (Thiel et al., 1995; Muro-Pastor et al., 1999). NtcA is a regulator required for expression of ammoniumrepressible genes; in a ntcA mutant, induction of nifHDK and hetR is abolished or minimal (Frias et al., 1994; Wei et al., 1994). The ntcA gene, which is conserved among cyanobacteria, bears a DNA-binding motif close to the C-terminus and is homologous to E. coli Crp and to S. meliloti FixK. The NtcA protein binds to defined sequence signatures that are located upstream of ammonium-regulated promoters (Luque et al., 1994). However, no such signature has been identified upstream of nif or hetR genes. The ntcA gene is autoregulated and presumed activators or cofactors may render NtcA active (MuroPastor et al., 1999). Biological N2 fixation requires a minimum of 16 ATP molecules and 8 reducing equivalents per molecule of N2 reduced. Under physiological conditions, a small electron carrier such as a ferredoxin or a flavodoxin is thought to transfer electrons to nitrogenase. In the photosynthetic

CHAPTER 1.24

Dinitrogen-Fixing Prokaryotes N-limitation

Utase (GlnD) PII

PII U NtrB P

NtrC (inactive)

NtrC (active) NtrB PII (GlnB)

807

activities are liberating huge amounts of fixed nitrogen to the environment (Socolow, 1999; Karl et al., 2002; McIsaac et al., 2002; Van Breemen et al., 2002), and as a consequence, nitrogen could become less limiting in nature and this may counterselect N2-fixing prokaryotes. Will some of them disappear without ever been known? After more than a century of research on N2 fixation, there are still ambitious goals to achieve. Acknowledgements My thanks to Julio Martínez Romero for technical help, and to Otto Geiger and Michael Dunn for reviewing the manuscript.

nifLA

Literature Cited

NifA

Austin, S., M. Buck, W. Cannon, T. Eydmann, and R. Dixon. 1994. Purification and in vitro activities of the native nitrogen fixation control proteins NifA and NifL. J. Bacteriol. 176:3460–3465. Achouak, W., P. Normand, and T. Heulin. 1999. Comparative phylogeny of rrs and nifH genes in the Bacillaceae. Int. J. Syst. Bacteriol. 49:961–967. Andrade, G., E. Esteban, L. Velasco, M. J. Lorite, and E. J. Bedmar. 1997. Isolation and identification of N2-fixing microorganism from the rhizosphere of Capparis spinosa (L.). Plant Soil 197:19–23. App, A. A., T. Santiago, C. Daez, C. Menguito, V. Ventura, A. Tirol, J. Po, et al. 1984. Estimation of the nitrogen balance for irrigated rice and the contribution of phototrophic nitrogen fixation. Field Crop Res. 9:17–27. App, A. A., I. Watanabe, T. S. Ventura, M. Bravo, and C. D. Jurey. 1986. The effect of cultivated and wild rice varieties on the nitrogen balance of flooded soil. Soil Sci. 141:448–452. Araujo da Silva, K. R., J. F. Salles, L. Seldin, and J. D. van Elsas. 2003. Application of a novel Paenibacillus-specific PCR-DGGE method and sequence analysis to assess the diversity of Paenibacillus spp. in the maize rhizosphere. J. Microbiol. Meth. 54:213–231. Arnold, W., A. Rump, W. Klipp, U. B. Priefer, and A. Pü hler. 1988. Nucleotide sequence of a 24,206-base-pair DNA fragment carrying the entire nitrogen fixation gene cluster of Klebsiella pneumoniae. J. Molec. Biol. 203:715– 738. Arsene, F., P. A. Kaminski, and C. Elmerich. 1996. Modulation of NifA activity by PII in Azospirillum brasilense: Evidence for a regulatory role of the NifA N-terminal domain. J. Bacteriol. 178:4830–4838. Awonaike, K. O., S. K. A. Danso, and F. Zapata. 1993. The use of a double isotope (15N and 34S) labelling technique to assess the suitability of various reference crops for estimating nitrogen fixation in Gliricidia sepium and Leucaena leucocephala. Plant Soil 155/156:325–328. Baker, D. D., and B. C. Mullin. 1992. Actinorhizal symbioses. In: G. Stacey, R. H. Burris, and H. J. Evans (Eds.) Biological Nitrogen Fixation. Chapman and Hall. New York, NY. 259–292. Baldani, V. L. D., and J. Dö bereiner. 1980. Host-plant specificity in the infection of cereals with Azospirillum spp. Soil Biol. Biochem. 12:433–439. Baldani, I., V. L. D. Baldani, L. Seldin, and J. Dö bereiner. 1986. Characterization of Herbaspirillum seropedicae

u-uridylylated p-phosphorylated Fig. 3. Cascade regulatory mechanisms in the g and b Proteobacteria under N-limited conditions. Uridylylated PII, as a cofactor of NtrB promotes the phosphorylation of NtrC that then becomes active to bind the upstream regulatory sequences (UAS) of the nifLA promoter. NifA in turn binds the UAS of the nitrogenase structural genes in many dinitrogen-fixing prokaryotes studied, allowing their expression and consequentely nitrogen fixation.

bacterium Rhodobacter capsulatus, a ferredoxin Fd1 was identified as the major electron donor to nitrogenase (Schatt et al., 1989; Schmehl et al., 1993). Conclusions Dinitrogen fixation is an important biological process carried out only by prokaryotes. Research on nitrogen fixation has followed a multidisciplinary approach that ranges from studies at the molecular level to practical agricultural applications. Support for research in this area has been driven by economic and environmental imperatives on the problems associated with the use of chemically synthesized nitrogen fertilizer in agriculture (Brewin and Legocki, 1996; Vance, 1998). However, the contributions of researchers in N2 fixation to gene regulation, biochemistry, physiology, microbial ecology, protein assembly, and structure, and more recently to genomics are highly meritorious achievements in themselves. Dinitrogen fixation research is a fast evolving field with specific model systems studied in great depth and an extensive knowledge of a larger diversity of N2-fixing prokaryotes more slowly developing. The advent of molecular biology has certainly enriched our knowledge of the reservoir of N2-fixing microorganisms and their ecology, but still the estimates of the amounts of nitrogen fixed in nature are uncertain. Human

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gen. nov., sp. nov., a root-associated nitrogen-fixing bacterium. Int. J. Syst. Bacteriol. 36:86–93. Baldani, J. I., V. M. Reis, V. L. D. Baldani, and J. Dö bereiner. 1999. Biological nitrogen fixation (BNF) in nonleguminous plants: The role of endophytic diazotrophs. In: F. O. Pedrosa, M. Hungria, et al. (Eds.) 12th International Congress on Nitrogen Fixation, Book of Abstracts. Universidade Federal do Paraná . Paraná,Brazil. 12. Bally, R., D. Thomas-Bauzon, T. Heulin, J. Balandreau, C. Richard, and J. De Ley. 1983. Determination of the most frequent N2-fixing bacteria in a rice rhizosphere. Can. J. Microbiol. 29:881–887. Barraquio, W. L., M. R. de Guzman, M. Barrion, and I. Watanabe. 1982. Population of aerobic heterotrophic nitrogen fixing bacteria associated with wetland and dryland rice. Appl. Environ. Microbiol. 43:124–128. Barraquio, W. L., J. K. Ladha, and I. Watanabe. 1983. Isolation and identification of N2-fixing Pseudomonas associated with wetland rice. Can. J. Microbiol. 29:867–873. Barraquio, W. L., L. Revilla, and J. K. Ladha. 1997. Isolation of endophytic diazotrophic bacteria from wetland rice. Plant Soil 194:15–24. Bauer, C. C., L. Scappino, and R. Haselkorn. 1993. Growth of the cyanobacterium Anabaena on molecular nitrogen: NifJ is required when iron is limited. Proc. Natl. Acad. Sci. USA 90:8812–8816. Berge, O., T. Heulin, W. Achouak, C. Richard, R. Bally, and J. Balandreau. 1991. Rahnella aquatilis, a nitrogen-fixing enteric bacterium associated with the rhizosphere of wheat and maize. Can. J. Microbiol. 37:195–203. Bergersen, F. J., and E. H. Hipsley. 1970. The presence of N2fixing bacteria in the intestines of man and animals. J. Gen. Microbiol. 60:61–65. Bergersen, F. J. 1974. Formation and function of bacteroids. In: A. Quispel (Ed.) The Biology of Nitrogen Fixation. North-Holland Publishing Company. Amsterdam, The Netherlands. 473–498. Bergersen, F. J. (Ed.). 1980. Methods for Evaluating Biological Nitrogen Fixation. Wiley. Chichester, UK. 701. Bergman, B., A. N. Rai, C. Johansson, and E. Sö derbäck. 1992. Cyanobacterial-plant symbioses. Symbiosis 14:61– 81. Berry, A. M. 1994. Recent developments in the actinorhizal symbioses. Plant Soil 161:135–145. Bianciotto, V., C. Bandi, D. Minerdi, M. Sironi, H. V. Tichy, and P. Bonfante. 1996. An obligately endosymbiotic mycorrhizal fungus itself harbors obligately intracellular bacteria. Appl. Environ. Microbiol. 62:3005–3010. Bianciotto, V., E. Lumini, P. Bonfante, and P. Vandamme. 2003. “Candidatus Glomeribacter gigasporarum” gen. nov., sp. nov., an endosymbiont of arbuscular mycorrhizal fungi. Int. J. Syst. Evol. Microbiol. 53:121–124. Bishop, P. E., and R. Premakumar. 1992. Alternative nitrogen fixation systems. In: G. Stacey, R. H. Burris, and H. J. Evans (Eds.) Biological Nitrogen Fixation. Chapman and Hall. New York, NY. 736–762. Boddey, R. M., S. Urquiaga, V. Reis, and J. Dö bereiner. 1991. Biological nitrogen fixation associated with sugar cane. Plant Soil 137:111–117. Boddey, R. M., O. C. de Oliveira, S. Urquiaga, V. M. Reis, F. L. de Olivares, V. L. D. Baldani, and J. Dö bereiner. 1995. Biological nitrogen fixation associated with sugar cane and rice: Contributions and prospects for improvement. Plant Soil 174:195–209. Bolin, J. T., A. E. Ronco, T. V. Morgan, L. E. Mortenson, and N.-H. Xuong. 1993. The unusual metal clusters of nitro-

CHAPTER 1.24 genase: Structural features revealed by x-ray anomalous diffraction studies of the MoFe protein from Clostridium pasteurianum. Proc. Natl. Acad. Sci. USA 90:1078– 1082. Bordeleau, L. M., and D. Prévost. 1994. Nodulation and nitrogen fixation in extreme environments. Plant Soil 161:115–125. Bottomley, P. J. 1992. Ecology of Bradyrhizobium and Gluconoacetobacter diazotrophicus obium. In: G. Stacey, R. H. Burris, and H. J. Evans (Eds.) Biological Nitrogen Fixation. Chapman and Hall. New York, NY. 293–348. Brewin, N. J., and A. B. Legocki. 1996. Biological nitrogen fixation for sustainable agriculture. Trends Microbiol. 4:476–477. Brigle, K. E., W. E. Newton, and D. R. Dean. 1985. Complete nucleotide sequence of the Azotobacter vinelandii nitrogenase structural gene cluster. Gene 37:37–44. Brigle, K. E., M. C. Weiss, W. E. Newton, and D. R. Dean. 1987. Products of the iron-molybdenum cofactor-specific biosynthetic genes, nifE and nifN, are structurally homologous to the products of the nitrogenase molybdenum-iron protein genes, nifD and nifK. J. Bacteriol. 169:1547–1553. Burgess, B. K., and D. J. Lowe. 1996. Mechanism of molybdenum nitrogenase. Chem. Rev. 96:2983–3012. Bü rgmann, H., F. Widmer, W. von Sigler, and J. Zeyer. 2003. mRNA extraction and reverse transcription-PCR protocol for detection of nifH gene expression by Azotobacter vinelandii in soil. Appl. Environ. Microbiol. 69:1928– 1935. Bü rgmann, H., F. Widmer, W. von Sigler, and J. Zeyer. 2004. New molecular screening tools for analysis of free-living diazotrophs in soil. Appl. Environ. Microbiol. 70:240– 247. Caballero-Mellado, J., and E. Martínez-Romero. 1994. Limited genetic diversity in the endophytic sugarcane bacterium Acetobacter diazotrophicus. Appl. Environ. Microbiol. 60:1532–1537. Caballero-Mellado, J., L. E. Fuentes-Ramírez, V. M. Reis, and E. Martínez-Romero. 1995. Genetic structure of Acetobacter diazotrophicus populations and identification of a new genetically distant group. Appl. Environ. Microbiol. 61:3008–3013. Caballero-Mellado, J., and E. Martínez-Romero. 1999. Soil fertilization limits the genetic diversity of Gluconoacetobacter diazotrophicus obium in bean nodules. Symbiosis 26:111–121. Carmichael, W. W. 1994. The toxins of cyanobacteria. Sci. Am. 270:64–72. Carrasco, C. D., J. A. Buettner, and J. W. Golden. 1995. Programed DNA rearrangement of a cyanobacterial hupL gene in heterocysts. Proc. Natl. Acad. Sci. USA 92:791– 795. Cavalcante, V. A., and J. Dö bereiner. 1988. A new acidtolerant nitrogen-fixing bacterium associated with sugarcane. Plant Soil 108:23–31. Cheetham, B. F., and M. E. Katz. 1995. A role for bacteriophages in the evolution and transfer of bacterial virulence determinants. Molec. Microbiol. 18:201–208. Chelius, M., and E. Triplett. 2001. The diversity of Archaea and Bacteria in association with the roots of Zea mays L. Microb. Ecol. 41:252–263. Chen, T.-H., S.-Y. Pen, and T.-C. Huang. 1993. Induction of nitrogen-fixing circadian rhythm Synechococcus RF-1 by light signals. Plant Sci. 92:179–182.

CHAPTER 1.24 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. Chen, W.-M., L. Moulin, C. Bontemps, P. Vandamme, G. Béna, and C. Boivin-Masson. 2003. Legume symbiotic nitrogen fixation by b-proteobacteria is widespread in nature. J. Bacteriol. 185:7266–7272. Christiansen-Weniger, C., A. F. Groneman, and J. A. van Veen. 1992. Associative N2 fixation and root exudation of organic acids from wheat cultivars of different aluminum tolerance. Plant Soil 139:167–174. Cojho, E. H., V. M. Reis, A. C. G. Schenberg, and J. Dö bereiner. 1993. Interactions of Acetobacter diazotrophicus with an amylolytic yeast in nitrogen-free batch culture. FEMS Microbiol. Lett. 106:341–346. David, M., M. L. Daveran, J. Batut, A. Dedieu, O. Domergue, J. Ghai, C. Hertig, P. Boistard, and D. Kahn. 1988. Cascade regulation of nif gene expression in R. meliloti. Cell 54:671–683. Dean, D. R., R. A. Setterquist, K. E. Brigle, D. J. Scott, N. F. Laird, and W. E. Newton. 1990. Evidence that conserved residues Cys-62 and Cys-154 within the Azotobacter vinelandii nitrogenase MoFe protein a-subunit are essential for nitrogenase activity but conserved residues His-83 and Cys-88 are not. Molec. Microbiol. 4:1505– 1512. Dean, D. R., and M. R. Jacobson. 1992. Biochemical genetics of nitrogenase. In: G. Stacey, R. H. Burris, and H. J. Evans (Eds.) Biological Nitrogen Fixation. Chapman and Hall. New York, NY. 763–834. DeLuca, T. H., O. Zackrisson, M. C. Nilsson, and A. Sellstedt. 2002. Quantifying nitrogen-fixation in feather moss carpets of boreal forests. Nature 419:917–920. D’hooghe, I., J. Michiels, K. Vlassak, C. Verreth, F. Waelkens, and J. Vanderleyden. 1995. Structural and functional analysis of the fixLJ genes of R. leguminosarum biovar phaseoli CNPAF512. Molec. Gen. Genet. 249:117–126. Dilworth, M. J., K. Fisher, C.-H. Kim, and W. E. Newton. 1998. Effects on substrate reduction of substitution of histidine-195 by glutamine in the a-subunit of the MoFe protein of Azotobacter vinelandii nitrogenase. Biochemistry 37:17495–17505. Distel, D. L., W. Morrill, N. MacLaren-Toussaint, D. Franks, and J. Waterbury. 2002. Teredinibacter turnerae gen. nov., sp. nov., a dinitrogen-fixing, cellulolytic, endosymbiotic gamma-proteobacterium isolated from the gills of wood-boring molluscs (Bivalvia: Teredinidae). Int. J. Syst. Evol. Microbiol. 52:2261–2269. Dixon, R., Q. Cheng, G.-F. Shen, A. Day, and M. DowsonDay. 1997. Nif gene transfer and expression in chloroplasts: Prospects and problems. Plant Soil 194:193–203. Dixon, R. 1998. The oxygen-responsive NIFL-NIFA complex: A novel two-component regulatory system controlling nitrogenase synthesis in g-proteobacteria. Arch. Microbiol. 169:371–380. Dixon, R. 1999. Prospects for engineering nitrogen-fixing photosynthetic eukaryotes. In: F. O. Pedrosa, M. Hungria, et al. (Eds.) 12th International Congress on Nitrogen Fixation, Book of Abstracts. Universidade Federal do Paraná . Paraná,Brazil. L034. Dö bereiner, J. 1961. Nitrogen-fixing bacteria of the genus Beijerinckia Derx in the rhizosphere of sugarcane. Plant Soil 15:211–217.

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into a phe-tRNA gene. Proc. Natl. Acad. Sci. USA 95:5145–5149. Sullivan, J. T., J. R. Trzebiatowski, R. W. Cruickshank, J. Gouzy, S. D. Brown, R. M. Elliot, D. J. Fleetwood, N. G. McCallum, U. Rossbach, G. S. Stuart, J. E. Weaver, R. J. Webby, F. J. de Bruijn, and C. W. Ronson. 2002. Comparative sequence analysis of the symbiosis island of Mesorhizobium loti strain R7A. J. Bacteriol. 184:3086– 3095. Suominen, L., M. M. Jussila, K. Makelainen, M. Romantschuk, and K. Lindstrom. 2000. Evaluation of the GalegaGluconoacetobacter diazotrophicus obium galegae system for the bioremediation of oil-contaminated soil. Environ. Pollut. 107:239–244. 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. Tan, Z., T. Hurek, and B. Reinhold-Hurek. 2003. Effect of N-fertilization, plant genotype and environmental conditions on nifH gene pools in roots of rice. Environ. Microbiol. 5:1009–1015. Thiel, T., E. M. Lyons, J. C. Erker, and A. Ernst. 1995. A second nitrogenase in vegetative cells of a heterocystforming cyanobacterium. Proc. Natl. Acad. Sci. USA 92:9358–9362. Trâ n Van, V., P. Mavingui, O. Berge, J. Balandreau, and T. Heulin. 1994. Promotion de croissance du riz inoculépar une bactérie fixatrice d’azote, Burkholderia vietnamiensis, isolée d’un sol sulfatéacide du Viet-nam. Agronomie 14:697–707. Trâ n Van, V., O. Berge, J. Balandreau, S. NgôKê, and T. Heulin. 1996. Isolement et activiténitrogénasique de Burkholderia vietnamiensis, bacterie fixatrice d’azote associée au riz (Oryza sativa L) cultivésur un sol sulfaté du Vietnam. Agronomie 16:479–491. Triplett, E. W. 1996. Diazotrophic endophytes: Progress and prospects for nitrogen fixation in monocots. Plant Soil 186:29–38. Turner, S. L., X. X. Zhang, F. D. Li, and J. P. Young. 2002. What does a bacterial genome sequence represent? Mis-assignment of MAFF 303099 to the genospecies Mesorhizobium loti. Microbiology (Reading, UK) 148:3330–3331. 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. Urquiaga, S., P. B. L. Botteon, and R. M. Boddey. 1989. Selection of sugar cane cultivars for associated biological nitrogen fixation using 15N-labelled soil. In: F. A. Skinner, et al. (Eds.) Nitrogen Fixation with Nonlegumes. Kluwer Academic Publishers. Dordrecht, The Netherlands. 311–319. Urquiaga, S., K. H. S. Cruz, and R. M. Boddey. 1992. Contribution of nitrogen fixation to sugar cane: Nitrogen-15 and nitrogen-balance estimates. Soil Sci. Soc. Am. J. 56:105–114. Vaisanen, O. M., A. Weber, A. Bennasar, F. A. Rainey, H. J. Busse, and M. S. Salkinoja-Salonen. 1998. Microbial communities of printing paper machines. J. Appl. Microbiol. 84:1069–1084. Valderrama, B., A. Davalos, L. Girard, E. Morett, and J. Mora. 1996. Regulatory proteins and cis-acting elements involved in the transcriptional control of Gluconoaceto-

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bacter diazotrophicus obium etli reiterated nifH genes. J. Bacteriol. 178:3119–3126. Van Breemen, N., E. Boyer, C. Goodale, N. Jaworski, K. Paustian, S. Seitzinger, K. Lajtha, B. Mayer, D. van Dam, R. Howarth, K. Nadelhoffer, M. Eve, and G. Billen. 2002. Where did all the nitrogen go? Fate of nitrogen inputs to large watersheds in the northeastern U.S.A. Biogeochemistry 57:267–293. Vance, C. P. 1998. Legume symbiotic nitrogen fixation: Agronomic aspects. In: H. P. Spaink, A. Kondorosi, and P. J. J. Hooykaas (Eds.) The Gluconoacetobacter diazotrophicus obiaceae. Kluwer Academic Publishers. Dordrecht, The Netherlands. 509–530. Vandamme, P., J. Goris, W.-M. Chen, P. De Vos, and A. Willems. 2002. Burkholderia tuberum sp. nov. and Burkholderia phymatum sp. nov., nodulate the roots of tropical legumes. Syst. Appl. Microbiol. 25:507– 512. Vaneechoutte, M., P. Kämpfer, T. De Baere, E. Falsen, and G. Verschraegen. 2004. Wautersia gen. nov., a novel genus accommodating the phylogenetic lineage including Ralstonia eutropha and related species, and proposal of Ralstonia [Pseudomonas] syzygii (Roberts et al. 1990) comb. nov. Int. J. Syst. Evol. Microbiol. 54:317–327. Ventura, T. S., M. Bravo, C. Daez, V. Ventura, I. Watanabe, and A. App. 1986. Effects of N-fertilizers, straw, and dry fallow on the nitrogen balance of a flooded soil planted with rice. Plant Soil 93:405–411. Vermeiren, H., W.-L. Hai, and J. Vanderleyden. 1998. Colonisation and nifH expression on rice roots by Alcaligenes faecalis A15. In: K. A. Malik, M. S. Mirza, and J. K. Ladha (Eds.) Nitrogen Fixation with Non-legumes. Kluwer Academic Publishers. Dordrecht, The Netherlands. 167–177. Vermeiren, H., A. Willems, G. Schoofs, R. de Mot, V. Keijers, W. Hai, and J. Vanderleyden. 1999. The rice inoculant strain Alcaligenes faecalis A15 is a nitrogen-fixing Pseudomonas stutzeri. System. Appl. Microbiol. 22:215– 224. Vlassak, K. M., and J. Vanderleyden. 1997. Factors influencing nodule occupancy by inoculant rhizobia. Crit. Rev. Plant Sci. 16:163–229. Von der Weid, I., G. F. Duarte, J. D. van Elsas, and L. Seldin. 2002. Paenibacillus brasilensis sp. nov., a novel nitrogenfixing species isolated from the maize rhizosphere in Brazil. Int. J. Syst. Evol. Microbiol. 52:2147–2153. Waelkens, F., A. Foglia, J.-B. Morel, J. Fourment, J. Batut, and P. Boistard. 1992. Molecular genetic analysis of the R. meliloti fixK promoter: Identification of sequences involved in positive and negative regulation. Molec. Microbiol. 6:1447–1456. Walcott, J. J., M. Chauviroj, A. Chinchest, P. Choticheuy, R. Ferraris, and B. W. Norman. 1977. Long term productivity of intensive rice cropping systems on the central plains of Thailand. Exp. Agric. 13:305–316. Wang, E. T., M. A. Rogel, A. Garcia-de los Santos, J. Martínez-Romero, M. A. Cevallos, and E. MartínezRomero. 1999. Rhizobium etli bv. mimosae, a novel biovar isolated from Mimosa affinis. Int. J. Syst. Bacteriol. 49:1479–1491. Wang, E. T., P. van Berkum, X. H. Sui, D. Beyene, W. X. Chen, and E. Martínez-Romero. 1999. Diversity of rhizobia associated with Amorpha fructicosa isolated from Chinese soils and description of Mesorhizobium amorphae sp. nov. Int. J. Syst. Bacteriol. 49:51–65.

CHAPTER 1.24 Wang, E. T., and E. Martínez-Romero. 2000a. Phylogeny of root- and stem-nodule bacteria associated with legumes. In: E. W. Triplett (Ed.) Prokaryotic Nitrogen Fixation. Horizon Scientific Press. Wymondham, UK. 177–186. Wang, E., and E. Martínez-Romero. 2000. Sesbania herbacea-Rhizobium huautlense nodulation in flooded soils and comparative characterization of S. herbaceanodulating rhizobia in different environments. Microb. Ecol. 40:25–32. Watanabe, I., R. So, J. K. Ladha, Y. Katayama-Fujimura, and H. Kuraishi. 1987. A new nitrogen-fixing species of pseudomonad: Pseudomonas diazotrophicus sp. nov. isolated from the root of wetland rice. Can. J. Microbiol. 33:670–678. Watanabe, I., T. Yoneyama, B. Padre, and J. K. Ladha. 1987. Difference in natural abundance of 15N in several rice (Oryza sativa L.) varieties: Applications for evaluating N2 fixation. Soil Sci. Plant Nutr. 33:407–415. Waters, J. K., B. L. Hughes, 2nd, L. C. Purcell, K. O. Gerhardt, T. P. Mawhinney, and D. W. Emerich. 1998. Alanine, not ammonia, is excreted from N2-fixing soybean nodule bacteroids. Proc. Natl. Acad. Sci. USA 95:12038– 12042. Webster, G., V. Jain, M. R. Davey, C. Gough, J. Vasse, J. Denarie, and E. C. Cocking. 1998. The flavonoid naringenin stimulates the intercellular colonization of wheat roots by Azorhizobium caulinodans. Plant Cell Environ. 21:373–383. Wei, T.-F., T. S. Ramasubramanian, and J. W. Golden. 1994. Anabaena sp. strain PCC 7120 ntcA gene required for growth on nitrate and heterocyst development. J. Bacteriol. 176:4473–4482. Welbaum, G. E., F. C. Meinzer, R. L. Grayson, and K. T. Thornham. 1992. Evidence for and consequences of a barrier to solute diffusion between the apoplast and vascular bundles in sugarcane stalk tissue. Australian J. Plant. Physiol. 19:611–623. Wernegreen, J. J., and M. A. Riley. 1999. Comparison of the evolutionary dynamics of symbiotic and housekeeping loci: A case for the genetic coherence of rhizobial lineages. Molec. Biol. Evol. 16:98–113. Wolk, C. P. 1996. Heterocyst formation. Ann. Rev. Genet. 30:59–78. Yamada, Y., K. Hoshino, and T. Ishikawa. 1997. The phylogeny of acetic acid bacteria based on the partial sequences of 16S ribosomal RNA: The elevation of the subgenus Gluconoacetobacter to the generic level. Biosci. Biotechnol. Biochem. 61:1244–1251. Yanni, Y. G., R. Y. Rizk, V. Corich, A. Squartini, et al. 1997. Natural endophytic association between R. leguminosarum bv. trifolii and rice roots and assessment of its potential to promote rice growth. Plant Soil 194:99– 114. Yoneyama, T., T. Muraoka, T. H. Kim, E. V. Dacanay, and Y. Nakanishi. 1997. The natural 15N abundance of sugarcane and neighbouring plants in Brazil, the Philippines and Miyako (Japan). Plant Soil 189:239–244. Young, J. P. W. 1992. Phylogenetic classification of nitrogenfixing organisms. In: G. Stacey, R. H. Burris, and H. J. Evans (Eds.) Biological Nitrogen Fixation. Chapman and Hall. New York, NY. 43–86. Youzhong, L., R. Parsons, D. A. Day, and F. J. Bergersen. 2002. Reassessment of major products of N sub(2) fixation by bacteroids from soybean root nodules. Microbiology 148:1959–1966.

CHAPTER 1.24 Zahran, H. H. 1999. Rhizobium-legume symbiosis and nitrogen fixation under severe conditions and in an arid climate. Microbiol. Molec. Biol. Rev. 63:968–989. Zanetti, S., U. A. Hartwig, A. Luescher, T. Hebeisen, M. Frehner, B. U. Fischer, G. R. Hendrey, H. Blum, and J. Noesberger. 1996. Stimulation of symbiotic N2 fixation in Trifolium repens L. under elevated atmospheric pCO2 in a grassland ecosystem. Plant Physiol. 112:575–583. Zehr, J. P., M. Mellon, S. Braun, W. Litaker, T. Steppe, and H. W. Paerl. 1995. Diversity of heterotrophic nitrogen fixation genes in a marine cyanobacterial mat. Appl. Environ. Microbiol. 61:2527–2532.

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Zehr, J. P., M. T. Mellon, and S. Zani. 1998. New nitrogenfixing microorganims detected in oligotrophic oceans by amplification of nitrogenase (nifH) genes. Appl. Environ. Microbiol. 64:3444–3450. Zehr, J. P., B. D. Jenkins, S. M. Short, and G. F. Steward. 2003. Nitrogenase gene diversity and microbial community structure: A cross-system comparison. Environ. Microbiol. 5:539–554. Zhang, Y., E. L. Pohlmann, P. W. Ludden, and G. P. Roberts. 2003. Regulation of nitrogen fixation by multiple PII homologs in the photosynthetic bacterium Rhodospirillum rubrum. Symbiosis 35:85–100.

Prokaryotes (2006) 2:818–841 DOI: 10.1007/0-387-30742-7_25

CHAPTER 1.25 tooR

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a i re t caB

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semugeL

Root and Stem Nodule Bacteria of Legumes MICHAEL J. SADOWSKY AND P. H. GRAHAM

Introduction The root and stem nodule bacteria of legumes (collectively called the rhizobia) comprise a genetically diverse group of organisms characterized by the ability to produce swellings or nodules on the stems or roots of most, but not all, leguminous plants (peas, beans, clover, etc.). Not all legumes form nodules with rhizobia. Within the nodules, rhizobia convert atmospheric dinitrogen (N2) gas into ammonia. This fixed nitrogen (N) is subsequently assimilated by the host, and improves plant growth and productivity. Approximately 300 million hectares (Mha) of legumes are grown worldwide, and they collectively fix about 60 tere grams (Tg) (6 ¥ 107 metric tons) of N each year (Kinzig and Socolow, 1994). Overall, N2 fixation supplies about 50% of the N used in agriculture, and because the fixed N is used directly by the host plant without initial passage through the soil, the process is generally considered environmentally friendly (Vance, 1998). Fixation rates vary with plant species, length of the growing season, presence of a suitable microsymbiont, and environmental conditions, but rates commonly are in the range of 100–200 kg of N2 fixed ha-1 yr-1 (Sparrow et al., 1995; Unkovich et al., 1997). Because of the practical benefits of nodulation and N2 fixation, the rhizobia have been extensively studied, particularly the genetic basis for their symbiotic interactions. However, the rhizobia are also good saprophytes, with soil populations of 103 to 104 rhizobia g-1 being common in soils previously used for legume growth. Thus, the ecological attributes of these organisms also have been studied extensively.

Phylogeny The taxonomy of the organisms producing root and stem nodules on legumes is in a state of flux. Though this ever-changing taxonomy affects what the organisms are called and how they are distinguished, it has little impact on their phylogenetic relationships. Small subunit

rRNA sequence analysis (SSU rRNA) supports division of these organisms into three major groups (Rhizobium [including Agrobacterium, Allorhizobium, Sinorhizobium, and Mesorhizobium], Bradyrhizobium and Azorhizobium) within the " subclass of the Proteobacteria (Martinez-Romero and Caballero-Mellado, 1996; Young and Haukka, 1996b). Wang et al., 1998 show that Bradyrhizobium and Azorhizobium are only distantly related to fast-growing Rhizobium and their relatives. Figure 1 also highlights divisions within Rhizobium that in the late 1980s through 1990s, led to subdivision of this genus as indicated above (Chen et al., 1988; DeLajudie et al., 1994; de Lajudie et al., 1998a; Jarvis et al., 1997). These changes, however, are currently under challenge (Kuykendall et al., 2000). Also notable in this figure is the overlap between species of Rhizobium and Agrobacterium. Amalgamation of Rhizobium and Agrobacterium has been proposed on a number of occasions (Graham, 1964; Heberlein et al., 1967; De Ley, 1968; Sawada et al., 1993; Parker, 1957), suggesting that the rhizobia may have evolved from plant pathogenic bacteria. Nonpathogenic Agrobacterium are well known as nodule contaminants (Hofer, 1941; Graham, 1976; de Lajudie et al., 1999), and often are confused with the nodule-forming rhizobia. Relative to the large number of species of Rhizobium that have been described, only a limited number of Bradyrhizobium and Azorhizobium species have been distinguished. This is likely to change as additional tropical legume species are studied. Additional groups of bradyrhizobia have already been identified, but not detailed phylogenetically (So et al., 1994; Graham et al., 1995). Moreover, links between Rhizobium and related root nodule bacteria (Phyllobacterium, Brucella, and Bartonella) and between Bradyrhizobium and Rhodopseudmonas, Blastobacter, and Afipia have been described, but need additional study. The ability to form nodules is restricted to a clade of plants including both legume and actinorhizal species. Not all legumes bear nodules, the percentage of plant species with nodules increasing from only 23% in the more primitive

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and changes in rhizobial taxonomy and could have included more than one species of rhizobia. As new legumes are commercialized and exploited, studies to examine the extent of legume/microsymbiont biodiversity near the legume’s center of origin, and to explore the consequences of founder effects are warranted.

Taxonomy

Fig. 1. Initiation of nodule formation on the roots of Phaseolus vulgaris (L.) by Rhizobium etli 8 days after inoculation. Photo by permission of M. H. Chaverra.

Caesalpinioideae to 97% in the Papilionoideae. Because these groups of legumes differ in the frequency of nodulation (and because Rhizobium, Bradyrhizobium, and Azorhizobium are so different), it has been suggested that the ability to nodulate and fix N2 could have arisen on more than one occasion (Parker, 1968; Doyle, 1994; Doyle and Doyle, 1997). Doyle, 1994 suggested that nodulation has arisen on at least three previous occasions, including in the genus Chamaecrista. Species within Chamaecrista can be distinguished from the non-nodulated, but closely related, Cassia on the basis of randomly amplified polymorphic DNA (RAPD) analysis (Whitty et al., 1994). Within Chamaecrista, species differ in the retention and release of rhizobia from infection threads during differentiation (Naisbitt et al., 1992). It seems unlikely that legumes as different as Phaseolus and Acacia could nodulate with both Rhizobium and Bradyrhizobium (Lange, 1961; Michiels et al., 1998). The rhizobia associated with a particular legume host can show significant diversity (Pinero et al., 1988; Souza et al., 1994). However, some caution in interpreting results from biodiversity studies is advisable because a number of studies predate recent phylogenetic advances

Rhizobia have traditionally been a difficult group to classify. Early researchers considered all rhizobia part of a single species that could nodulate any legume. Subsequently each rhizobial strain was shown to only nodulate certain specific legumes. This led to the concept of cross inoculation groups, with rhizobia being distinguished according to the legumes each could nodulate. Thus, rhizobia from alfalfa would generally nodulate medic species and vice versa, but neither would nodulate clover. Using this approach, more than 20 different cross-inoculation groups were identified, and a number of these were raised to species status within the Rhizobium (Fred et al., 1932). Fred et al. (1932) stated, “It seems true that the ability of an organism to infect certain plants and not others is as fixed and definite as any phase of the physiology of the organism . . . we feel justified in regarding it as the prime character in species differentiation.” Host specificity is still important in the identification of rhizobia but is often at odds with results from numerical and phylogenetic studies (Graham, 1964; DeLey and Rassell, 1965; Heberlein et al., 1967; Moffett and Colwell, 1968). The demonstration that the nodulation genes in Rhizobium may be plasmid borne (Nuti et al., 1979; Brewin et al., 1980) or located on chromosomal symbiotic islands and move between organisms has further weakened infection-based taxonomic analyses. The 1984 edition of Bergey’s Manual of Systematic Bacteriology divides the Rhizobiaceae into four groups, including three genera of nodule- or gall-forming bacteria, Rhizobium, Bradyrhizobium and Agrobacterium (Jordan, 1984). The reduced emphasis on host range and the availability of several new phenotypic and phylogenetic techniques has resulted in the proliferation of new species and genera of nodule bacteria. Currently, 6 genera and 28 species of rhizobia are recognized (Table 1). Kuykendall et al. (2000) question the need for separation of Rhizobium, Agrobacterium, Allorhizobium and Sinorhizobium, and instead suggest the consolidation of these organisms into a single genus Rhizobium, having three subgenera. In the classification they propose, undicola, galegae and huautlense are included in the subgenus

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CHAPTER 1.25 Table 1. Listing of validly published species of root and stem nodule bacteria. Species Rhizobium 1. R. leguminosarum bvs. trifolii, viciae and phaseoli 2. R. galegae 3. R. tropici 4. R. etli 5. R. gallicum 6. R. giardini 7. R. huautlense 8. R. mongolense Sinorhizobium 1. S. meliloti 2. S. fredii 3. S. saheli 4. S. teranga 5. S. medicae Mesorhizobium 1. M. loti 2. M. huakuii 3. M. ciceri 4. M. tianshanense

Fig. 2. A Bradyrhizobium japonicum cell showing significant polyhydroxybutyrate accumulation. Photo by T. McDermott, used with permission.

Agrobacterium, with three species of plant pathogenic agrobacteria. With new species of root-nodule bacteria now justified using a polyphasic approach that includes both phenotypic and phylogenetic traits (Graham et al., 1991), the further description of new species of rhizobia based solely on simple characteristics has become increasingly problematic. In the second edition of The Prokaryotes, (Elkan and Bunn, 1994) listed phenotypic traits useful in the distinction of Rhizobium, Azorhizobium and Bradyrhizobium. To do so again is a daunting and perhaps not a useful task because older species descriptions may include more than one organism, and because differences in the tests applied and methods used can impact results and their interpretation. Table 2 lists carbon source utilization differences for many of the species of root and stem nodule bacteria. It was compiled from a number of different studies (Jarvis et al., 1997; de Lajudie et al., 1998b; Wang et al., 1999) and will likely change as new species are identified. Additional distinctive phenotypic differences are urgently needed. Analysis of rhizobial fatty acid methyl esters (FAME), using gas chromatography (Jarvis and

5. M. mediterraneum 6. M. plurifarium 7. M. amorphae Allorhizobium 1. Al. undicola Brudyrhizobium 1. B. elkanii 2. B. joponicum 3. B. liaoningense Azorhizobium 1. Az. caulinodans

Reference Frank, 1989 Frank, 1989 Lindstrom, 1989 Martinez et al., 1991 Segovia et al., 1993 Amarger et al., 1997 Amarger et al., 1997 Wang et al., 1998 van Berkum et al., 1998 de Lajudie et al., 1994 Dangeard, 1926 de Lajudie et al., 1994 Scholla and Elkan, 1984 de Lajudie et al., 1994 de Lajudie et al., 1994 de Lajudie et al., 1994 Rome et al., 1996 Jarvis et al., 1997 Jarvis et al., 1982, 1997 Chen et al., 1991 Jarvis et al., 1997 Nour et al., 1994 Jarvis et al., 1997 Chen et al., 1995 Jarvis et al., 1997 Nour et al., 1995 Jarvis et al., 1997 de Lajudie et al., 1998 Wang et al., 1999 de Lajudie et al., 1998 de Lajudie et al., 1998 Jordan, 1982 Kuykendall et al., 1992 Kirchner, 1895 Jordan, 1982 Xu et al., 1995 Dreyfus et al., 1998 Dreyfus et al., 1988

Tighe, 1994; Jarvis et al., 1996), has been recommended as a relatively simple and inexpensive method for the identification of fast-growing rhizobia. Rhizobial FAME profiles correctly identified nearly 95% of almost 200 strains evaluated by (Jarvis and Tighe, 1994 and Jarvis et al., 1996). These studies only erred in identifying some fredii as meliloti and etli as leguminosarum and vice versa (Graham et al., 1999). Ballen and Graham (unpublished observations) have also shown that etli, gallicum, and strains from Dalea and Onobrychisoverlap. Similarly, FAME profiles have been used to distinguish slow-growing japonicum and elkanii (Kuykendall et al., 1992; So et al., 1994; Graham et al., 1995), though in each case additional isolates were identified that did not group with these species.

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Table 2. Differences among genera of root nodule bacteria in the carbon compounds used for growth 1. Genus of nodule bacteria Carbon source Adonitol D-Arabinose L-Arabinose D-Cellobiose L-Fucose Inositol Gluconate Lactose L-Lysine DL-Malate D-Maltose D-Mannose Mannitol D-Mellibiose D-Raffinose Ribose L-Rhamnose Sucrose Trehalose D-Xylose

Rhizobium

Sinorhizobium

Mesorhizobium

Allorhizobium

+ +

+ +

+ + + + +

+ +/+ (+) + +/(+) + + (+) + + + + + + +

+ + (+)

+ + + + + + + + + + (-) (-) -

(+) + + + + + + + + + +

+/+/(+) (-) +/+ (+) + (-) +/+ + + +

Bradyrhizobium + + + (+) + (+) + (+) + (+) +

Azorhizobium + + -

Symbols: +, positive reaction; -, negative reaction; +/-, discriminatory within the genus; (+), mainly positive reaction; (-), mainly negative reaction. 1 Includes data from Elkan and Brunn, 1992; de Lajudie et al., 1994, 1998; Rome et al., 1996; Jarvis et al., 1997 and Wang et al., 1999.

Habitat Rhizobia through their ability to fix N2 in symbiosis with legumes play a central role in the N supply of most natural ecosystems. The American tall grass prairie is but one ecosystem in which plant diversity and productivity is controlled in large measure by N availability (Collins et al., 1998). Rhizobia, although thought to be solely soil saprophytes, can also be found in aquatic systems associated with water-growing leguminous plants. Owing to cultural and agricultural practices, the migration of birds and animals, and atmospheric deposition of soil particles, there are relatively few soils in the world that do not contain some rhizobia. Rhizobia have been shown to exist in soils for a relatively long time in the absence of a host plant (Bottomley, 1992; Brunel et al., 1988; Kucey and Hynes, 1989; Sanginga et al., 1994; Slattery and Coventry, 1993; Weaver et al., 1972). Rhizobia have been recognized as being important for the functioning of soil ecosystems for centuries (Fred et al., 1932). Shortly, after legume root nodules were shown conclusively to assimilate atmospheric N2 (Hellriegel and Wilfarth, 1888), Nodbe and Hiltner applied for, and were granted, a patent for the use of these microorganisms as legume inoculants (Elkan and Bunn, 1994). This and subsequent farming and cultural practices have led to the dissemination of rhizobia on a global basis.

Rhizobia in soils may be introduced by application of commercial inoculants or, as in many cases, be the normal flora present as microsymbionts of an indigenous legume. Inoculants applied to seed, as recommended by their manufacturer, achieve inoculation rates of 103–106 rhizobia seed-1 (Somasegaran and Hoben, 1994). This corresponds to application rates of up to 8 ¥ 1010 rhizobia ha-1 (Brockwell and Bottomley, 1995). At these rates, inoculant strains often dominate in nodulation in the first year of a newly introduced crop (Brockwell et al., 1982; Gibson et al., 1976; Singleton and Tavares, 1986). Moreover, inoculant strains contribute to the rapid buildup of rhizobia in the soil once nodulessenesce and release large numbers of viable rhizobia into the soil system (McDermott et al., 1987; Sutton, 1983). Several studies have documented that inoculant strains dominate in nodules 5–15 years after initial inoculation (Brunel et al., 1988; Diatloff, 1977; Lindstrom et al., 1990). It should be noted, however, that not all introduced legumes receive inoculation, and in such situations, seed, soil or aerial contamination will usually lead to some initial nodule formation, and over a period of 4–5 years, a buildup of soil rhizobial populations (Sadowsky and Graham, 1998a). Moreover, diverse rhizobial populations can develop in association with species that are not initially indigenous to a particular region (Leung et al., 1994). Although it is thought that rhizobia in soil have a clonal

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population structure, genetic recombination between groups of soil rhizobia may be contributing to diversity in soils (Demezas et al., 1995; Sullivan et al., 1995). It has been demonstrated that soil rhizobia can transfer plasmids (Jarvis et al., 1989; Kinkle and Schmidt, 1991; Thomas et al., 1994; Young and Wexler, 1988) and chromosomal symbiotic genes (Sullivan et al., 1995). The rhizobia obtained from any given soil habitat are drastically influenced by the common method of isolation. This usually involves the use of serial dilutions of soil and inoculation on a trap host, followed by recovery from nodules (Somasegaran and Hoben, 1994). This procedure often underestimates the numbers of rhizobia in the soil and biases diversity determinations (Dye et al., 1995). Numerous studies have documented the influence of a trap host on the recovery of particular groups of rhizobia from soils (Bottomley et al., 1994; Bromfield et al., 1995; Brunel et al., 1996; Keatinge et al., 1995; Kumar Rao et al., 1982; van Berkum et al., 1995). Selective culture media, when available, will most likely prove useful in determining the identity of natural populations of rhizobia in soil (Gault and Schwinghamer, 1993; Tong and Sadowsky, 1993). Lastly, the legume host itself has been shown to strongly influence the prevalence and type of rhizobia in soils (Bottomley, 1992). How this occurs is not known, but it is thought to be due to nonspecific, root-exudate enhanced growth of rhizobia in the rhizosphere, multiplication and release of rhizobia from the nodule, and selection by the trap host of particular groups of rhizobia from mixed soil populations (Sadowsky and Graham, 1998a).

Isolation Date (1982); Date and Halliday (1987) and Somasegaran and Hoben (1994) have detailed methods for the collection, sampling, isolation, authentication and maintenance of rhizobia. Extensive collection and conservation is necessary because many isolates will prove to be ineffective in symbiosis, or host/strain interactions will be significant. In the case of Stylosanthusscabia, more than 1,000 isolates were evaluated before a strain suitable for use in commercial legume inoculation was identified (Date, 1997).

Collection The collection of rhizobia is most commonly undertaken as part of a plant introduction program, to supply suitable host germplasm with the rhizobia they need for symbiosis (see the National Plant Germplasm Collection System http://www.ars-grin.gov). Ideally, the collection

CHAPTER 1.25

of nodules should coincide with early season growth and well watered conditions. However, where collection involves remote geographic regions, sample acquisition may be delayed until plant maturity when most nodules may have senesced. Nodule collection may also be limited where the plant species in question is endangered, and no plant harvest is permitted. In both of these cases, soil may be used as a source of rhizobia, using surface-sterilized seed of an appropriate host to “trap” nodule bacteria. Collection of rhizobia also may be undertaken to study the biodiversity of indigenous organisms, or to study success in nodulation, or the soil establishment of bioengineered organisms. In some cases, the culture of rhizobia from nodules may be unnecessary because enough cell material may be present in soils or plant tissue to directly characterize nodule occupants using serological or phylogenetic methods (Sadowsky and Graham, 1998). Somasegaran and Hoben (1994) list several culture collections of rhizobia throughout the world. In addition, the USDAARS (National Rhizobium Resource Collectionl) (bldg6.arsusda.gov) provides a searchable database of rhizobia grouped by legume host.

Sampling Sampling of plants and nodules should be done from undisturbed locations and, where possible, from healthy plants. Accurate site description and record keeping are essential. The number of nodules needed can vary with the reason for collection. Where the aim is to identify inoculantquality rhizobia, 15–20 nodules per plant, taken from the crown region of the host root system, are usually sufficient. Where the goal is to evaluate strain biodiversity in soil, a large number of nodules should be collected from as much of the root system as possible. Ease of collection may vary; stoloniferous species may have nodules on adventitious roots within 1–2 cm of the surface (Date, 1982), while nodules on tree species may be at a great depth in the soil at some distance from the trunk of the tree. Collected nodules should be protected in vacutainers or in vials containing a dessicant (e.g., silica gel) overlain by cotton wool (Somasegaran and Hoben, 1994).

Isolation Successful isolation of rhizobia from nodules depends on the quality of nodules recovered. When nodules have been stored dry over silica gel or CaCl2, they must first be allowed to imbibe (sterile) water fully before being surface sterilized. Rhizobia also can be frequently recovered from nodules obtained from intact root system frozen at -20∞C. Sodium hypochlorite (3%),

CHAPTER 1.25

Root and Stem Nodule Bacteria of Legumes

hydrogen peroxide (3%) and acidified mercuric chloride (0.1%) are all effective surface sterilants. The former is usually preferred due to its low cost, ready availability and ease of disposal. Surface sterilization procedures are described in detail by Vincent (1970) and Somasegaran and Hoben (1994). Yeast extract mannitol (YEM) medium is commonly used in the routine isolation and subculture of of rhizobia. Many different formulations for this medium exist (Vincent, 1970; Somasegaran and Hoben, 1994). That used in our laboratory contains: Mannitol MgSO4 ·7H 2O NaCl K2HPO4 CaCl2 ·2H 2O FeCl3 ·6H 2O Yeast extract Agar Distilled water pH

10.0 g 0.2 g 0.1 g 0.5 g 0.2 g 0.01 g 1.0 g 20.0 g 1 liter 6.7–7.0

Sterilize by autoclaving 20 min at 103 H 103 pascal (15 lb/in2) pressure. The medium may be amended with cycloheximide (20 microgram/ml) to reduce fungal contamination and bromthymol blue (BTB; 25 microgram/ml) or Congo red (25 microgram/ml) to facilitate identification of rhizobia. These should be filter-sterilized separately and added to autoclaved, molten YEM medium before plates are poured (Vincent, 1970). Tong and Sadowsky (1993) described a selective medium specific for Bradyrhizobium, based primarily on the heavy metal tolerance of these organisms. Media selective for fast-growing rhizobia have been described (Barber, 1979; Louvrier et al., 1995), but they have not proven generally effective. Rhizobium, Mesorhizobium, Sinorhizobium, and Allorhizobium strains will generally produce moist, gummy colonies on YEM medium that are 4–6 mm in diameter after 7 days incubation. On medium containing BTB, the colonies and surrounding medium are yellow due to acid production by the microorganisms. Slower growing bradyrhizobia produce smaller colonies, usually only 1–2 mm diameter after 7–10 days incubation, which are raised and mucoid. The colonies and surrounding medum are blue in color on YEM containing BTB. Most nodule isolates will produce white or cream colored colonies, though some isolates produce melanin (Cubo et al., 1988), or in the case of bradyrhizobia, a rust red pigmentation in older colonies.

Authentication Authentication of rhizobia usually involves completion of Koch’s postulates with the host from

823

which strains were originally isolated. Somasegaran and Hoben (1994) provide details of this methodology. Inoculated seedlings produced from surface-sterilized seed of a suitable legume host are typically grown in sterile low N medium or on seedling agar in large test tubes, growth pouches or Leonard jars. Plants are examined for nodulation after 25–30 days of incubation under lights. The presence of nodules on uninoculated control plants invalidates the experiment.

Identification The identity of rhizobia or bradyrhizobia often requires a multiphasic approach using many of the techniques employed in naming new genera, species and strains of rhizobia (Graham et al., 1991). Members of the International Subcommittee on Rhizobium and Agrobacterium, a subcommittee of the International Committee on Systematic Bacteriology of the International Union of Microbiological Societies, have recommended a minimal set of criteria for naming new species and genera of nodule bacteria (Graham et al., 1995). These criteria are also useful for identifying the genus and species status of unknown rhizobia isolated from nodules. In addition to biochemical, cultural, and symbiotic data, 16S rRNA (rDNA) sequencing (Young, 1996a; Young and Haukka, 1996b), DNA-DNA hybridization (Scholla et al., 1984), FAME (Graham et al., 1995), and multilocus enzyme electrophoresis, (MLEE) (Strain et al., 1995) data are of primary importance for identifying rhizobia isolated from newly surveyed legumes. Following strain authentication, it is often useful to mark these isolates to facilitate identification in subsequent ecological, genetic or plant studies. This can be done using a variety of techniques. These include intrinsic resistance to a series of different antibiotics (Josey et al., 1979) or the selection of mutants resistant to high levels of antibiotics. In the latter case, selected mutants must be evaluated to show that the acquisition of antibiotic resistance has not influenced nodulation, N2 fixation or competitive abilities. Strains can also be identified using strain or group-specific antibodies (Sadowsky et al., 1987a; Schmidt et al., 1968). Antibodies, which are typically produced in rabbits to somatic whole-cell antigens, are useful in strain identification because they do not require genetic modification of strains. Agglutination, immunodiffusion, immunoflourescence, and ELISA techniques all have found wide acceptance in serological identification of rhizobia (Dudman, 1977; Humphrey and Vincent, 1965; Kishinevsky and Jones, 1987; Schmidt et al., 1968). The fluo-

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rescent antibody technique is especially useful because it allows for the direct in situ examination of rhizobia in soil and nodules (Bohlool and Schmidt, 1973), using direct or sandwich labeling procedures. Strains also can be genetically modified with b$-glucuronidase reporter (GUS) (Wilson et al., 1995; Wilson et al., 1999) and lux (Chabot et al., 1996) genes, and these strains have proven especially useful in ecological studies. Again, however, it is essential that such genetically marked strains be plant tested before use in ecological, symbiotic, or field studies. DNA fingerprinting techniques have been used to identify and study biodiversity of rhizobial strains (Sadowsky, 1994; Versalovic et al., 1998; Sadowsky and Hur, 1998b; Demezas, 1998). Initially, DNA fingerprints of strains were generated following restriction enzyme digestion of total genomic DNA (Glynn et al., 1985; Demezas, 1998). More recently, however, restriction fragment polymorphism (RFLP) analysis techniques, DNA hybridization probes, and DNA primers corresponding to repetitive elements, coupled to the polymerase chain reaction (PCR) technique, have been used in strain identification, competition, and biodiversity studies (de Bruijn, 1992; Judd et al., 1993; Sadowsky, 1994; Sadowsky et al., 1990; Wheatcroft and Watson, 1988).

Cultivation Rhizobia are relatively robust, ubiquitous, aerobic bacteria with the ability to utilize many different substrates (carbon [C] and nitrogen [N] sources) for growth (Parke and Ornston, 1984). Consequently, rhizobia can be cultivated on a large variety of complex and defined culture media. Only a limited number of rhizobia grow on highly enriched media, such as nutrient broth or LB medium. Medium used in the cultivation of rhizobia depends on the species of nodule bacteria, growth characteristics desired, and the method of cultivation. Most rhizobia are mesophiles and can grow in shake cultures at 25–30∞C. However, rhizobia isolated from legumes grown in the Canadian High Arctic grow well at 5∞C (Prevost et al., 1987), and high temperature tolerant strains have been isolated in Africa and Brazil. As stated earlier, most rhizobia grow well in YEM medium (Vincent, 1970), though most produce copious quantities of capsular- and exo-polysaccharides in this medium, limiting its use in biochemical and genetic studies. The bradyrhizobia grow fairly slowly in this medium, with generation times greater than 6 hours. Rhizobia and bradyrhizbia shift the pH of this

CHAPTER 1.25

medium, the rhizobia produce acid and the bradyrhizobia, alkaline byproducts, from growth. Polysaccharide production can be drastically reduced in fast-growing rhizobia by cultivation in TY medium (Beringer, 1978), containing (g/liter): Tryptone (5.0), Yeast extract (3.0) and CaCl2 ·2H 2O (0.87), pH 6.9. In this medium turbid cultures, up to 109 cells ml-1, can be obtained after overnight incubation at 28∞C. The slowgrowing bradyrhizobia do not grow in TY medium. A growth medium useful for polysaccharide-free growth by bradyrhizobia is AG medium (Sadowsky et al., 1987b). This medium, which promotes rapid growth of japonicum and elkanii strains, contains (g/liter): HEPES (0.13), MES (0.11), FeCl3 ·6H 2O (0.0067), MgSO4 ·7H 2O (0.18), CaCl2 ·2H 2O (0.013), Na2SO4 (0.25), NH4Cl (0.32), Na2HPO4 (0.125), arabinose (1.0), Na-gluconate (1.0) and yeast extract (1.0), pH 6.9. Several defined, minimal, media are also used for the growth of rhizobia (Vincent, 1970; Somasegaran and Hoben, 1994), especially for biochemical and molecular biological studies. We also use AG medium without arabinose, gluconate, and yeast extract, as a minimal medium for the cultivation of prototrophic rhizobia and in genetic mating studies. Rhizobia for legume inoculants can be grown in shake flasks or fermentors (Somasegaran and Hoben, 1994). Lorda and Balatti (1996) described a glycerol-based culture medium capable of producing approximately 1010 cells/ml, even in shake flask culture. In contrast, Stephens and Rask (2000) suggest that carbon-limited media be used to produce legume inoculants, to condition rhizobia to the less favorable conditions found in soil.

Preservation Rhizobium strains in frequent use are usually maintained on YMA slants in screw capped test tubes stored at 6–10∞C. Longer-term storage is achieved by lyophilization with 10% glycerol or 10% sucrose and 5% peptone as cryoprotectants, or by storage at -70∞C in 15% glycerol. Gherna (1994) details methodologies for lyophilization and storage at -70∞C. Change in Rhizobium characteristics with repeated growth on laboratory media has been reported (Herridge and Roughley, 1975) and must be of concern. Some inoculant companies maintain large numbers of ampoules of each Rhizobium strain in the freezedried state Rhizobium and routinely replace all working cultures at 3-month to 1-year intervals.

The Nodulation Process The nodulation process requires molecular communication between both symbiotic part-

CHAPTER 1.25

ners and involves the induction and repression of a large number of bacterial and plant genes. Free-living rhizobia infect and form N2fixing symbioses with legumes in a series of discrete stages or steps. Stages in the process include: proliferation of rhizobia in the rhizosphere, recognition of host by rhizobia, attachment of rhizobia to susceptible root hair cells, root-hair curling and infection-thread formation, initiation of nodule primordium, and transformation of free-living rhizobia into N2fixing bacteroids. Rhizobia infect their respective host plants and induce root or stem nodules using several different mechanisms. Infection through root hairs is commonly seen with most legumes (Hadri et al., 1998). Rhizobia can also invade the host plant by entry through wounds, cracks, or lesions caused by emergence of secondary roots (Boogerd and van Rossum, 1997), as occurs in peanut and Stylosanthes. In these cases, rhizobia spread intercellularly. There are instances where the same rhizobia infect one legume through root hairs and another via cracks or wounds (Sen and Weaver, 1988). Lastly, rhizobia may initiate infection of the host via cavities surrounding adventitious root primordia on the stems of Sesbania, Aeschynomene, Neptunia, and Discolobium (Boivin et al., 1997). As above, one bacterium may produce both stem and root nodules on different legume plants. Nodule shape in legumes is determined by the host plant and is regulated by the pattern of cortical cell divisions. There are two basic types of nodules that are formed on legumes: determinant and indeterminant (Franssen et al., 1992). Indeterminant nodules are most commonly formed in symbioses between the fast-growing nodule bacteria and temperate legumes (pea, clover, and alfalfa). Determinate nodules, which are normally induced by bradyrhizobia, are more common on tropical legumes, such as soybean and bean. Morphologically, indeterminate nodules have defined, persistent apical meristems and are elongated and sometimes lobed, whereas determinant nodules do not have persistent meristems and are usually round (Hadri et al., 1998). In root hair infection, rhizobia attach to susceptible root hairs within minutes of inoculation or contact with the host plant. Rhizobial cells often attach perpendicular to the root hair cell. It has been suggested that adhesion is initially mediated by the calcium (Ca)-binding protein rhicadhesin, or by plant lectins, and subsequent bonding via production of cellulose fibrils (Kijne, 1992). It is hypothesized that rhizobia produce localized hydrolysis of the root hair cell wall. Subsequent penetration of rhizobia through the cell wall leads to root-hair curling, which may be

Root and Stem Nodule Bacteria of Legumes

825

visible 6–18 hours after inoculation. The proportion of root hairs infected is low, the percentage of these giving rise to nodules is low and highly variable, and aborted root hairs can frequently be found. Within the root hair, rhizobia are enclosed within a plant-derived infection thread, and move down the root hair in the direction of the root cortex. Cell division in the root cortex, in advance of the approaching infection thread, leads to the production of nodule primordia (Kijne, 1992). Spread of the infection thread among cells of the nodule primordium follows, with the release of rhizobia into host cortex by an endocytotic process. Rhizobia are never free in the cytoplasm, but rather are surrounded by a host-derived peribacteroid membrane, which serves to compartmentalize the rhizobia into a symbiosome. One to several rhizobia can be confined to a single symbiosome. Nodulation is usually visible 6–18 days after inoculation, but this varies considerably with the selection of bacterial strain and host cultivar, the inoculant density and placement, and the temperature. Initially nodulation is heaviest in the crown of the root, with secondary nodules appearing on lateral roots as the first-formed nodules senesce. The number of nodules produced on each legume host is tightly controlled by the host and rhizobial genotype, the efficiency of the symbiotic interaction, by environmental factors such as soil N level and the presence of existing nodules (Caetano-Annoles, 1997; Sagan and Gresshoff, 1996; Singleton and Stockinger, 1983).

Genetics The genetics of the rhizobia has, in most cases, centered on the genetics of nodulation and symbiotic N2-fixation, key characters that set the rhizobia apart from other soil bacteria. Recent advances in molecular biology and genetics have elucidated a large number of genes with symbiotic functions. Though many of these genes are clustered together (on the chromosome in some organisms and on symbiotic plasmids in others), additional genes may be dispersed or located on different replicons. Consequently, all symbiotically related genes will most likely not be found until total sequencing and functional genomic efforts are completed. Because the scope of this chapter is broad, more detailed information on the genetics of nodulation and N2 fixation can be found in several recent reviews (Boivin et al., 1997; van der Drift et al., 1998; Schultze and Kondorosi, 1998; Niner and Hirsch, 1998; Denarie et al., 1996; Pueppke, 1996; Spaink, 1995).

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Nodulation Genes In the last several years, a large number of bacterial genes have been identified which are involved in the formation of nodules on leguminous plants. Collectively, more than 65 nodulation genes have been identified in rhizobia, although each strain may only have a subset of these. Niner and Hirsch (1998); Pueppke (1996); and Bladergroen and Spaink (1998) provide a more complete description of the function of a majority of these genes. Several studies have shown that relatively few genes are required for nodulation of legumes (Gö ttfert, 1993; Long et al., 1985; Long, 1989; van Rhijn and Vanderleyden, 1995). In the case of the fast-growing rhizobia, a majority of nodulation genes are located on large, indigenous, symbiotic (Sym), and often self-transmissible, plasmids (Broughton et al., 1984; Hombrecher et al., 1981; Kondorosi et al., 1989). The complete genomic sequence of the symbiotic plasmid from Rhizobium sp. strain NGR-234, a Rhizobium strain with broad nodulation ability (Pueppke and Broughton, 1999), is currently available. In meliloti, the symbiont of alfalfa, nodulation genes (located on an 8.5 kb fragment of the Sym plasmid) contain sequences necessary for the nodulation of a wide variety of legume hosts (Kondorosi et al., 1989; Truchet et al., 1991). These genes, referred to as “common nodulation” genes and designated nodA, nodB and nodC, have homologues in other fast- and slowgrowing species. In leguminosarum bvs. trifolii and viceae and meliloti, the common nodulation genes are organized in a similar cluster (Downie et al., 1985; Egelhoff and Long, 1985; Fisher et al., 1985; Nieuwkoop et al., 1987; Putnoky and Kondorosi, 1986; Rolfe et al., 1985; Russell et al., 1985; Schofield and Watson, 1986; van Rhijn and Vanderleyden, 1995). A fourth gene, nodD, is regulatory and together with plant flavonoid signals (see below) activates transcription of other inducible nod genes (Long, 1989; Martinez et. al, 1990; van Brussel et al., 1990). Leguminosarm bvs. viceae and trifolii have single copies of nodD. The symbionts meliloti and japonicum have multiple copies of nodD (Gö ttfert et al., 1986; Gö ttfert et al., 1990; Honma and Ausubel, 1987). In some instances, nodD also appears to impart host-specificity functions (Spaink et al., 1987). Another nodulation gene cluster, originally designated hsn (for host-specific nodulation), is closely linked to the common nodulation region in meliloti and controls nodulation of specific legume genera (Bachem et al., 1986; Horvath et al., 1986). Mutations in the hsn genes (designated nodFEGH) cannot be complemented with Sym plasmids from other species of Rhizobium. Analogous hsn genes also have

CHAPTER 1.25

been isolated from leguminosarum bv. trifolii (Djordjevic et al., 1985; Rolfe et al., 1985), leguminosarum bv. viceae (Wijffelman et al., 1985), and from Rhizobium strain MPIK3030 (Bachem et al., 1986; Bassam et al., 1986; Broughton et al., 1984; Lewin et al., 1987). An hsn gene linked to the common nodulation region in japonicum strain USDA 110 has also been reported (Nieuwkoop et al., 1987). This sequence, subsequently called nodZ (Dockendorff et al.,1994), was shown to be involved in the host-specific nodulation of siratro, but not soybean. Hahn and Hennecke (1988) and Gö ttfert et al. (1990) have identified another hsn locus in japonicum strain 110, nodVW, which is essential for the nodulation of siratro, mungbean and cowpea, but not soybean. In japonicum strain USDA 110, the essential nodulation genes are located on the chromosome in several transcriptional units in the order: nolZ, nolA, nodD2, nodD1, nodYABCSUIJmolMNO (Dockendorff et al., 1994). Unlike other rhizobial nodD genes, the japonicum nodD1 is induced by the flavonoids genistein and daidzein (Banfalvi et al., 1988; Kosslak et al., 1987) and by xanthones (Zaat et al., 1987).

Genotype-Specific Nodulation Genes Although many hsn genes have been identified in Rhizobium and Bradyrhizobium, there are only limited reports on the identification of genotype-specific nodulation (GSN) genes in the rhizobia (Sadowsky et al., 1991). The GSN genes specifically refer to those bacterial genes that allow nodulation of specific plant genotypes within a given legume species. For example, strain TOM nodulates the pea genotype Pisum sativum cv. Afghanistan (Lie, 1978a; Lie et al., 1978b), but European leguminosarum bv. viceae strains fail to nodulate this host. Some GSN-like genes have been found on plasmid pRL5JI of strain TOM (Gotz et al., 1985; Hombrecher et al., 1984). Davis et al. (1988) have identified a single gene on this plasmid, nodX, mediating the O-acetylation of Nod factors (Firmin et al., 1993), which is necessary for the nodulation of “Afghanistan” peas. In fredii strain USDA 257, two other GSN-like loci, nolC (Krishnan and Pueppke, 1991) and nolBTUVW (Meinhardt et al., 1993), allow this strain to nodulate primitive lines of soybean, but not improved soybean varieties, such as “McCall” (Heron et al., 1989). In each case, Tn5 insertions in the gene regions allow fredii to nodulate commercial soybean cultivars. Phenotypically, these regions are similar to that reported by Djordjevic et al. (1985) and Innes et al. (1985) for clover rhizobia. More recently, however, Lewis-Henderson and Djord-

CHAPTER 1.25

jevic (1991a) reported that nodM in leguminosarum bv. trifolii is a GSN which prevents effective nodulation of subterraneum cv. Woogenellup (Lewis-Henderson and Djordjevic, 1991b). Analysis of leguminosarum bv. trifolii strain TA1 demonstrated that this strain also lacks nodT, and that introduction of nodT from leguminosarum bv. viceae strain ANU843 into TA1 allows effective nodulation of “Woogenellup” (LewisHenderson Djordjevic, 1991a). The GSN genes can act in either a positive or negative manner (Djordjevic et al., 1987a; Sadowsky et al., 1990), insertions in a negatively acting nodulation gene extending host-range and insertions in a positively acting GSN gene limiting host range. Bradyrhizobium japonicum serogroup 123 strains are restricted for nodulation by PI 377578 (Cregan and Keyser, 1986). The japonicum nolA gene, identified in strain USDA 110, is a positively acting gene that allows serogroup 123 strains to nodulate PI 377578 (Sadowsky et al., 1991). We recently identified a mutant of japonicum strain USDA 110 that has the ability to overcome nodulation restriction conditioned by soybean PI 417566 (Lohrke et al., 1995).

Signal Exchange and Induction of Nod Genes Although the regulation of nodulation genes in rhizobia is still not fully understood, we know a lot about communication between rhizobia and susceptible legume hosts. Flavonoid signal molecules present in root and seed exudates are necessary for nod gene expression (Banfalvi et al., 1988; Boundy-Mills et al., 1994; Djordjevic et al., 1987a; Fellay et al., 1995; Gö ttfert et al., 1988; Innes et al., 1985; Kosslak et al., 1987; Long, 1989; Mulligan and Long, 1985; Olson et al., 1985; Peters et al., 1986; Price et al., 1992; Sadowsky et al., 1988; van Brussel et al., 1990; Zaat et al., 1987). Other Sym plasmid-borne genes are also induced by root exudates in fredii and Rhizobium sp. strain NGR234 (Boundy-Mills et al., 1994; Fellay et al., 1995; Olson et al., 1985; Sadowsky et al., 1988). Flavones, isoflavones, flavanols, flavanones, and closely related compounds have been identified as nod gene inducers, and each is specific for a particular legume-Rhizobium interaction (Schlaman et al., 1998). Flavanoid compounds are only one of several determinants of host specificity. Spaink et al. (1991) reported differential induction of nodD in various fast-growing rhizobia by a range of flavonoids and exudates. Induction of nodulation genes requires the regulatory nodD gene product (Long, 1989; Mulligan and Long, 1985; Shearman et al., 1986). The inducer apparently binds NodD, causing a change in conformation (Kondorosi et al., 1988; Fisher and Long, 1989).

Root and Stem Nodule Bacteria of Legumes

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Activated NodD then binds to a regulatory, promoter-like sequence, found upstream of rhizobial nod genes, the nod box (Hong et al., 1987; Horvath et al., 1986; Kondorosi et al., 1988; Rostas et al., 1986; Shearman et al., 1986). Repressor proteins have also been suggested to play a role in nod gene regulation (Kondorosi et al., 1988), a repressor encoded by the nolR gene has been identified in S. meliloti strain 41 (Kondorosi et al., 1989; Kondorosi et al., 1991).

Extracellular Nodulation Factors One of the primary functions of nod genes is the production of extracellular lipochitinoligosaccharide (LCO) molecules, also known as Nod factors (Carlson et al., 1993; Carlson et al., 1994). These molecules, acting at 10-8 to 10-9 M, can: 1) stimulate the plant to produce more nod gene inducers (van Brussel et al., 1990); 2) deform root hairs on homologous hosts (Banfalvi et al., 1989; Faucher et al., 1989); and 3) initiate cell division in the root cortex (Lerouge et al., 1990; Price et al., 1992; Sanjuan et al., 1992; Schultze et al., 1992; Relic et al., 1993; Spaink et al., 1991). In meliloti these signal molecules are acetylated and sulfated glucosamine oligosaccharides (Lerouge et al., 1990). Similar molecules have been identified in other legume symbiotic systems (Pueppke, 1996 and Downie, 1998 for a review). Numerous observations support the theory that hsn genes control host specificity by decorating Nod factors with various substituents. For example, the meliloti genes nodP, nodQ and nodH are involved in the sulfation of the Nod factor reducing sugar (Faucher et al., 1989; Roche et al., 1991). Disruption of any of these genes affects host specificity. Rhizobium spp. NGR234, which can nodulate over 125 different legume species (Pueppke and Broughton, 1999), produces diverse (more than 18) Nod factors, which vary in the substituents attached to a similar backbone structure (Price et al., 1992). Purified Nod factors, which are structurally similar to those produced by the appropriate rhizobial symbiont, can induce nodules on the specific host plant in the absence of a bacterium (Downie, 1998; Mergaert et al., 1993; Relic et al., 1993, Schultze et al., 1992; Truchet et al., 1991). Nod factors from several strains of japonicum have been characterized (Carlson et al., 1993; Sanjuan et al., 1992). The functions of nod genes and the basic structure of Nod factors for japonicum and several species of the genus Rhizobium can be found in (Downie, 1998).

Nitrogen Fixation Genes Two major types of N2 fixation genes have been described, nif genes and fix genes. The nif refer

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CHAPTER 1.25

to genes involved in the N2 fixation process and have structurally and functionally related genes in the free-living diazotrophic microorganism, Klebsiella pneumoniae. pneumoniae was the first N2-fixing microorganism studied in detail (Kennedy, 1989). As with the nodulation genes, a majority of the nif genes are plasmid borne and contiguous in the rhizobia, but chromosomally located in the bradyrhizobia. The N2 fixation process is catalyzed by the enzyme complex nitrogenase, encoded by the nifDK and nifH genes. The fix genes are also involved in the N2 fixation process, but have no similar structural or functional homologues in pneumoniae. The organization of nif genes varies in the rhizobia (Kaminski et al., 1998). Nitrogenase consists of two protein subunits, a molybdenum-iron (MoFe) protein and an ironcontaining (Fe) protein. These structural components of the nitrogenase enzyme complex are often referred to as subunits I and II, respectively. The nifK and nifD genes encode the MoFe protein subunits. A FeMo cofactor (FeMo-Co) is required for activation of the MoFe protein and is assembled from the nifB, V, N, and E genes. The nifH gene encodes the Fe subunit protein. In pneumoniae there are at least 20 nif-specific genes that are localized in about 8 operons (Dean and Jacobson, 1992). Though the organization of nif genes in other organisms varies tremendously (Downie, 1998), nifHD and nifK are conserved in disparate N2-fixing organisms and rhizobia (Ruvkin and Ausubel, 1980). The gene products NifA and NifL control the regulation of all other nif genes. Whereas NifA is positive activator of transcription of nif operons, NifL is involved in negative control. In K. pneumoniae and several other free-living diazotrophic microbes, nif gene expression is regulated by oxygen and nitrogen levels (Merrick, 1992). Ammonia (NH3) causes NifL to act as a negative control and prevents the activator function of NifA. This has been referred to as the N control system, and has been shown to regulate several enzymes that are capable of producing NH3. Merrick (1992) and Dean and Jacobson (1992)

give excellent in-depth reviews of the structure and regulation of N2 fixation in free-living and symbiotic bacteria.

Other Genes Involved in Symbiotic Nitrogen Fixation Other plasmid and chromosomally borne bacterial genes also have been found to function indirectly in nodulation and symbiotic N2-fixation in rhozobia (Table 3). Recent review articles on the structure and function of these and other symbiosis-related genes are provided by Pueppke et al. (1996) and Spaink (1995) and Long (1999).

Ecology Rhizobia are relatively unique among the majority of soil microorganisms in that they have an extensive soil phase as free-living, saprophytic, heterotrophic microorganisms, yet in conjunction with leguminous plants, they have the ability to form species-specific, N2-fixing symbiotic associations. The ability to form N2-fixing nodules affords unique opportunities for the rhizobia. When a legume crop is grown in soil for the first time, few rhizobia are likely to be present and, in most instances, inoculation will most likely be needed for adequate nodulation and subsequent N2 fixation (Date, 1991; Diatloff, 1977). In contrast, soils surrounding legumes that have been planted for several years usually contain relatively large numbers of rhizobia and do not require added rhizobia. Numerous studies have documented that legume inoculants added to soils containing relatively small populations of rhizobia usually give rise to only a small percentage of the nodules formed (Thies et al., 1991; Date, 1991; Ellis et al., 1984; Ham, 1978). Despite intensive investigations over the last 30 years, however, some of the factors that influence the survival and the persistence of rhizobia in the soil, their ecology and competitiveness for nodulation sites on the host, are only now beginning to be understood. It is beyond the scope of this

Table 3. Some other bacterial genes involved in symbiotic nitrogen fixation. Gene designation

Phenotype or function

Exo

Exopolysaccharide

Hup Gln Dct

Hydrogen uptake Glutamine synthase Dicarboxylate transport

Nfe Ndv LPS

Nodulation efficiency $-1,2 Glucans Lipopolysaccharide

Reference Bechei and Phhler, 1988 Glazebrook and Walker, 1989 Maier, 1986 Carlson et al., 1987 Finan et al., 1983 Jiang et al., 1989 Sanjuan and Olivares, 1989 Breedveld and Miller, 1998 Carlson et al., 1987

CHAPTER 1.25

chapter to present all that is known about the ecology and soil biology of the rhizobia. The reader is directed to more extensive reviews by Bottomley (1992) and Sadowsky and Graham (1998a) on this material. The establishment of the symbiotic state results in the production of a nodule populations of more than 1010 rhizobia g-1 nodule tissue (McDermott et al., 1987). When these nodules senesce at the end of the growing season, large numbers of rhizobia are released into the soil. Nodule bacteroids are subject to changes in surface chemistry (Roest et al., 1995) and are susceptible to osmotic and other soil stresses (Sutton, 1983). However, many of the released organisms manage to persist as free-living, heterotrophic, saprophytes in the soil until a susceptible legume is again planted. As a consequence of this, most soils contain at least some rhizobia, and a dramatic buildup in their numbers occurs when a leguminous host is included in a crop rotation, pasture or natural setting. Ellis et al. (1984) reported soil populations of bradyrhizobia approaching 106 cells g-1 in soils of the American Midwest following cultivation of soybean, and rhizosphere populations can reach 108 cells g-1 (Bottomley, 1992). The distribution of rhizobia in soil is not uniform. Postma et al. (1990) reported that the greatest number of rhizobia are associated with soil aggregates of larger than 50 mm, and Mendes and Bottomley (1998) noted that the percentage of Rhizobium recovered from aggregates of different sizes varied over the course of a growing season. Rhizobia are excellent soil saprophytes and can persist for many years in the absence of their host (Brunel et al., 1988; Kucey and Hynes, 1989; Bottomley, 1992). Chatel et al. (1968) used the term saprophytic competence to describe this ability, but the factors involved have yet to be determined. Even though Bushby, 1990 noted surface electrophoretic charge in bradyrhizobia correlated to the pH of soils from which they came, Rynne et al., 1994 found no correlation between catabolic ability and strain persistence. Inoculant strains used at the time a particular host was introduced may still occupy a large percentage of the nodules formed on that host 10– 15 years after their introduction (Diatloff, 1977; Brunel et al., 1988; Lindstrom et al., 1990). However, many studies have shown that inoculant strains may also decline in nodule representation over time, or quite quickly disappear from soil. The growth of rhizobia in the rhizosphere may also be stimulated by specific root exudates (Van Egaraat, 1975). Rhizobia, in turn, also stimulate growth and respiration of leguminous plants (Phillips et al., 1999). Several research studies have sought to create biased rhizospheres, in which plants transformed to synthesize opines,

Root and Stem Nodule Bacteria of Legumes

829

favor the growth of rhizosphere bacteria utilizing this substrate (Rossbach et al., 1994; Oger et al., 1997; Savka and Farrand, 1997). The inability of strains to compete for nodulation sites on the host legume does not necessarily mean their displacement from the soil population. Bromfield et al. (1995) compared populations of meliloti recovered from soil and nodules and found significant differences in the frequency with which particular genotypes were recovered. Similarly, Segovia et al. (1991) found the population of noninfective bean rhizobia in soil numerically superior to those capable of inducing nodule formation. Environmental factors, particularly soil pH, temperature and water availability, often affect rhizobial survival in soil, and the balance between particular genotypes. In soils of pH >7.0, Brockwell et al., 1991 found an average of 89,000 meliloti g-1 soil, whereas in soils of pH optimal

balanced redox state I O2 Fe3+

direction of motility parallel to magnetic field direction of motility ontiparalldl to magnetic field

Fe2+ S2–

reduced state [Red]in > optimal

balanced redox state II

[S2–]

Fig. 6. Hypothetical model of the function of polar magnetotaxis in bacteria (Northern hemisphere). Cells are guided along the geomagnetic field lines depending on their “redoxstate” either downward to the sulfide-rich zone or upward to the microoxic zone, thereby enabling a shuttling between different redox layers.

microcolony-like aggregates, fits well with this model, which is summarized in the following Fig. 6.

Morphologic and Phylogenetic Diversity Morphotypes The diversity of magnetotactic bacteria is reflected by the high number of different morphotypes found in environmental samples of water or sediment. Commonly observed morphotypes include coccoid to ovoid cells, rods, vibrios and spirilla of various dimensions. One of the more unique morphotypes is an apparently multicellular bacterium referred to as the MMP many-celled magnetotactic prokaryote. Regardless of their morphology, all magnetotactic bacteria studied so far are motile by means of flagella and have a cell wall structure characteristic of Gram-negative bacteria. The arrangement of flagella differs and can be either polar, bipolar, or in tufts. Another trait which shows considerable diversity is the arrangement of magnetosomes inside the bacterial cell. In the majority of magnetotactic bacteria, the magnetosomes are aligned in chains of various lengths and numbers along the cell’s long axis of the cell, which is magnetically the most efficient orientation. However, dispersed aggregates or clusters of magnetosomes occur in some magnetotactic bacteria usually at one side of the cell, which often corresponds to the site of flagellar insertion. Besides magnetosomes, large inclusion bodies containing elemental sulfur, polyphos-

847

phate, or poly-b-hydroxybutyrate are common in magnetotactic bacteria collected from the natural environment and in pure culture. The most abundant type of magnetotactic bacteria occurring in environmental samples, especially sediments, are coccoid cells possessing two flagellar bundles on one somewhat flattened side. This bilophotrichous type of flagellation gave rise to the tentative genus “Bilophococcus” for these bacteria (Moench, 1988). One representative strain of this morphotype is in axenic culture (see Other Magnetotactic Strains in Pure Culture in this Chapter). In contrast, two of the morphologically more conspicuous magnetotactic bacteria, regularly observed in natural samples but never isolated in pure culture, are the MMP and a large rod containing large numbers of hook-shaped magnetosomes (“Candidatus Magnetobacterium bavaricum”). The MMP, a Many-Celled Magnetotactic Prokaryote A magnetotactic aggregation of cells that swims as an entire unit and not as separate cells was first reported and described by Farina et al. (1983). Similar morphotypes were later found also in sulfide-rich marine and brackish waters and in sediments along the coasts of North America and Europe (Mann et al., 1990a). The MMP (for many-celled magnetotactic prokaryote) consists of about 10 to 30 coccoid to ovoid Gram-negative cells, roughly arranged in a sphere with a diameter ranging from approximately 3 to 12 mm (Fig. 7). Cells are asymmetrically multiflagellated on their outer surfaces exposed to the external surroundings. Magnetosomes consist of the magnetic iron-sulfide greigite, Fe3S4, and several nonmagnetic precursors to greigite (see “IronSulfide Type Magnetosomes”). The magnetosome crystals are generally pleomorphic although cubo-octahedral, rectangular prismatic, and tooth-shaped particles have also been observed in cells. They are usually loosely arranged in short chains or clusters in individual cells. The total magnetic moment of the MMP was determined and ranges from 5 ¥ 10-16 to 1 ¥ 10-15 Am2, which is sufficient for an effective magnetotactic response. The type of magnetotaxis displayed by the MMP appears to be polar, but aggregates have been observed to reverse direction. Under oxic conditions in a uniform magnetic field, the swimming speed in the preferred direction averages 105 mm/s. After reaching the edge of a water drop, aggregates sometimes spontaneously reverse their swimming direction and show short excursions of 100 to 500 mm with twice the speed of the forward motion in the opposite direction of their polarity (Rodgers et al., 1990) as shown in Fig. 8. This so

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CHAPTER 1.26

5 µm

a

b

1 µm

Fig. 7. Brightfield TEM micrographs of the many-celled magnetotactic prokaryote (MMP). (a) An unstained, single MMP revealing the numerous greigite-containing magnetosomes within the organism mostly arranged in short chains. (b) Negatively-stained preparation (2.5% ammonium molybdate, pH 7.0) of a single MMP that is disrupted to reveal separated individual cells. (c) Thin-section of an MMP again showing its many-celled nature. (d) Negatively-stained individual cell of the MMP. Note the asymmetric distribution of flagella which cover the cell on one side, the pleomorphism of the greigite-containing magnetosomes, and the electron-lucent vacuoles resembling poly-b-hydroxybutyrate (PHB) granules.

0.5 µm

c

d

called “ping pong” motion seems to be a peculiarity of this organism. Fig. 8. Sequence showing the typical “ping-pong” motility of the MMP. For the video, see the online version of The Prokaryotes.

In one study, it was reported that individual cells within the aggregate are connected by intercellular membrane junctions (Rodgers et al., 1990a; 1990b). However, the cohesive force among individual cells seems to be relatively weak because a lowering of the osmotic pressure leads to an immediate disruption of the aggregate into single nonmotile cells. “Candidatus Magnetobacterium bavaricum” The first phenotypic description of this morphotype by Vali et al. (1987) was based on cells collected from material retrieved from the littoral sediments of a large freshwater lake in Southern Germany (Lake Chiemsee). Later, similar bacteria were also found in sediments of other freshwater habitats in Germany and Brazil. After the determination of its phylogenetic relationship (Spring et al., 1993), this organism was given the candidatus status due to its unusual phenotypic traits which distinguish it from all other magnetotactic bacteria. “Magnetobacterium bavaricum” displays polar magnetotaxis and is

0.5 µm

preferentially found in the microoxic zone of sediments, although significant numbers are also found in anoxic regions of their habitat (Fig. 1). In situ hybridizations using a specific fluorescently-labeled oligonucleotide probe targeting the 16S rRNA of this organism enabled the detection of microcolonies of this bacterium on microscope slides immersed into sediment for several weeks (Fig. 9). Thus, there appears to be a tendency for “Magnetobacterium bavaricum” to adhere to particles located in microsites with preferred environmental conditions. Cells of “M. bavaricum” are large rods having dimensions of 1–1.5 x 6–9 mm and are motile by a polar tuft of flagella. The most impressive trait of this bacterium is the extremely high number of magnetosomes per cell. A single cell may contain up to a thousand hook-shaped magnetosomes usually arranged in 3–5 rope-shaped bundles oriented parallel to the long axis of the cell (Fig. 10). The magnetosomes consist of magnetite (Fe3O4) and have a length of 110–150 nm. The average total magnetic moment per cell was experimentally determined to be approximately 3 ¥ 10-14 Am2, which is about an order of magnitude higher than that of most other magnetotactic bacteria. The presence of large sulfur inclusions is typical for this bacterium and seems

CHAPTER 1.26

Magnetotactic Bacteria

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a

1 µm

a

100 µm

100 nm

b b

Fig. 10. Brightfield transmission electron microscope (TEM) micrographs of “M. bavaricum.” (a) Whole cell displaying bundles of magnetosome chains and sulfur globules. (b) Hook-shaped magnetite-type magnetosomes. Courtesy of M. Hanzlik.

to be dependent on environmental conditions. In an unidirectional magnetic field, cells swim forward (i.e., northward in the Northern Hemisphere) with an average speed of 40 mm/s with the flagella wound around the rotating cell. Gradients of some chemical substances lead to a reversal of the sense of flagellar rotation resulting in a swimming in the opposite direction for a short time.

c Fig. 9. In situ hybridization of a microscope slide grown over with sediment bacteria using fluorescently labeled oligonucleotide probes. (a) Phase contrast micrograph. (b+c) Same field viewed with epifluorescence microscopy enabling the detection of a specific probe binding to a signature region of the 16S rRNA of “M. bavaricum” (b), and of a probe with broad specifity hybridizing with the 16S rRNA of most known bacteria (c).

Composition and Structure of Magnetosome Crystals The magnetosome mineral phase in magnetotactic bacteria are tens-of-nanometer-sized crystals of an iron oxide and/or an iron sulfide. The mineral composition of the magnetosome is specific enough for it to be likely under genetic control, in that cells of several cultured magnetiteproducing magnetotactic bacteria still synthesize an iron oxide and not an iron sulfide, even when hydrogen sulfide is present in the growth medium

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CHAPTER 1.26 a

b

(100)

(111)

(111)

(111)

(111)

(111)

(011)

(010)

(111) c

d 100 110

111 101

011

(001)

(111) (111)

(110) 0.5 µm

101

011 110 (111) (010)

(100) Fig. 11. Unusually large magnetite crystals identified in coccoid magnetotactic bacteria retrieved from a lagoon near Rio de Janeiro, Brazil. Small black dots represent gold-labeled antibodies detecting a specifically bound polynucleotide probe complementary to a highly variable region of the 16S rRNA of these cells.

(211)

001 (011) e

(100)

f (100) (111)

(111)

(Meldrum et al., 1993a; Meldrum et al., 1993b). The size of the magnetosome mineral crystals also appears to be under control of the organism because the large majority of magnetotactic bacteria contain crystals displaying only a very narrow size range, from about 35 to 120 nm (Frankel et al., 1998). Magnetite and greigite particles in this range are stable single magnetic domains (Butler and Banerjee, 1975; Diaz-Rizzi and Kirschvink, 1992). Smaller particles would be superparamagnetic at ambient temperature and would not have stable, remanent magnetization. Larger particles would tend to form multiple domains, reducing the remanent magnetization. However, in some uncultured bacteria from the Southern Hemisphere exceptionally large magnetite-magnetosomes have been observed in some uncultured bacteria from the Southern hemisphere (Fig. 11), having dimensions well above the theoretically determined size limits of single domain magnetite (Spring et al., 1998). It remains unclear if the crystals in these bacteria are still of single-domain size or are multi-domain particles and why such unusually large crystals are formed by certain bacteria, but, interestingly, it seems that the crystal-size corresponds with the size and/or the growth phase of these bacteria, i.e., large cells possess larger crystals than smaller cells of the same type. In contrast to chemically synthesized magnetite and greigite crystals, biologically produced magnetosome mineral particles display a range of well-defined morphologies which can be classified as distinct idealized types (Fig. 12).

(111)

(101)

(010)

(111) (010) (001)

Fig. 12. Idealized magnetite (a–d) and greigite (e–f) crystal morphologies derived from high resolution TEM studies of magnetosome crystals from magnetotactic bacteria: (a) and (e) cubo-octahedrons; (b), (c), and (f) variations of pseudohexagonal prisms; (d) elongated cubo-octahedron. Numbers within parentheses refer to the faces of the crystal lattice planes on the surface of the crystal. Figure adapted from Heywood et al. (1991) and Mann and Frankel (1989).

The consistent narrow size range (Devouard et al., 1998) and morphologies of the intracellular magnetosome particles represent typical features of a biologically controlled mineralization and are clear indications that the magnetotactic bacteria exert a high degree of control over the biomineralization processes involved in magnetosome synthesis. Magnetite-Type Magnetosomes The iron oxidetype magnetosomes consist solely of magnetite, Fe3O4. The particle morphology of the magnetite crystals in magnetotactic bacteria varies but is extraordinarily consistent within cells of a single bacterial species or strain (Bazylinski et al., 1994). Three general morphologies of magnetite particles have been observed in magnetotactic bacteria using transmission electron microscopy (TEM; Blakemore et al., 1989; Mann et al.,

CHAPTER 1.26

a

b

c

Magnetotactic Bacteria

100 nm

100 nm

100 nm

Fig. 13. Morphologies of intracellular magnetite (Fe3O4) particles produced by magnetotactic bacteria. (a) Darkfield scanning-transmission electron microscope (STEM) image of a chain of cubo-octahedra in cells of an unidentified rodshaped bacterium collected from the Pettaquamscutt Estuary, Rhode Island, USA, viewed along a [111] zone axis for which the particle projections appear hexagonal. (b) Brightfield TEM image of a chain of prismatic crystals within a cell of strain MV-2, a marine vibrio, with parallelepipedal projections. (c) Brightfield TEM image of tooth-shaped (anisotropic) magnetosomes from an unidentified rodshaped bacterium collected from the Pettaquamscutt Estuary.

851

1990a; Stolz, 1993; Bazylinski et al., 1994). They include: 1) roughly cuboidal (Balkwill et al., 1980; Mann et al., 1984); 2) parallelepipedal (rectangular in the horizontal plane of projection; Moench and Konetzka, 1978; Towe and Moench, 1981; Moench, 1988; Bazylinski et al., 1988); and 3) tooth-, bullet-, or arrowheadshaped (anisotropic; Mann et al., 1987a; Mann et al., 1987b; Thornhill et al., 1994). High resolution TEM and selected area electron diffraction studies have revealed that the magnetite particles within magnetotactic bacteria are of relatively high structural perfection and have been used to determine their idealized morphologies (Matsuda et al., 1983; Mann et al., 1984a; 1984b; 1987a; 1987b; Meldrum et al., 1993a; Meldrum et al., 1993b). These morphologies are all derived from combinations of {111}, {110} and {100} forms (a form refers to the equivalent symmetry related lattice planes of the crystal structure) with suitable distortions (Devouard et al., 1998). The roughly cuboidal particles are cubo-octahedra ([100] + [111]), and the parallelepipedal particles are either pseudohexahedral or pseudooctahedral prisms. Examples are shown in Fig. 12a–d. The cubo-octahedral crystal morphology preserves the symmetry of the face-centered cubic spinel structure, i.e., all equivalent crystal faces develop equally. The pseudohexahedral and pseudo-octahedral prismatic particles represent anisotropic growth in which equivalent faces develop unequally (Mann and Frankel, 1989; Devouard et al., 1998). The synthesis of the tooth-, bullet- and arrowheadshaped magnetite particles (Figs. 10b, 13c) appears to be more complex than that of the other forms. They have been examined by high resolution TEM in one uncultured organism (Mann et al., 1987a; Mann et al., 1987b) and their idealized morphology suggests that growth of these particles occurs in two stages. The nascent crystals are cubo-octahedra which subsequently elongate along the [111] axis parallel to the chain direction. Whereas the cubo-octahedral form of magnetite can occur in inorganically-formed magnetites (Palache, 1944), the prevalence of elongated pseudohexahedral or pseudo-octahedral habits in magnetosome crystals imply anisotropic growth conditions, e.g., a temperature gradient, a chemical concentration gradient, or an anisotropic ion flux (Mann and Frankel, 1989). This aspect of magnetosome particle morphology has been used to distinguish magnetosome magnetite from detrital or magnetite produced by biologically induced mineralization (by the anaerobic iron-reducing bacteria), using electron microscopy of magnetic extracts from sediments (e.g., Petersen et al., 1986; Chang and Kirschvink, 1989a; Chang et al., 1989b; Stolz et al., 1986; Stolz et al., 1990; Stolz, 1993).

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Iron-Sulfide Type Magnetosomes Virtually all freshwater magnetotactic bacteria have been found to synthesize magnetite as the mineral phase of their magnetosomes. In contrast, many marine, estuarine, and salt marsh species produce iron sulfide-type magnetosomes consisting primarily of the magnetic iron sulfide, greigite, Fe3S4 (Heywood et al., 1990; Heywood et al., 1991; Mann et al., 1990b; Pó sfai et al., 1998a; 1998b). Reports of non-magnetic iron pyrite (FeS2; Mann et al., 1990b) and magnetic pyrrhotite (Fe7S8; Farina et al., 1990) have not been confirmed and may represent misidentifications of additional iron sulfide species occasionally observed with greigite in cells (Pó sfai et al., 1998a; Pó sfai et al., 1998b). Currently recognized greigite-producing magnetotactic bacteria includes the MMP (Farina et al., 1983; Rodgers et al., 1990a; 1990b; DeLong et al., 1993) and a variety of relatively large, rod-shaped bacteria (Bazylinski et al., 1990; Bazylinski et al., 1993a; Heywood et al., 1990; Heywood et al., 1991; Bazylinski and Frankel, 1992). The iron sulfide-type magnetosomes contain either particles of greigite (Heywood et al., 1990; Heywood et al., 1991) or a mixture of greigite and transient non-magnetic iron sulfide phases that appear to represent mineral precursors to greigite (Pó sfai et al., 1998a; Pó sfai et al., 1998b). These phases include mackinawite (tetragonal FeS) and possibly a sphalerite-type cubic FeS (Pó sfai et al., 1998a; Pó sfai et al., 1998b). Based on TEM observations, electron diffraction, and known iron sulfide chemistry (Berner, 1967; Berner, 1970; Berner, 1974), the reaction scheme for greigite formation in the magnetotactic bacteria appears to be: cubic FeS Æ mackinawite sfai et al., (tetragonal FeS) Æ greigite (Fe3S4; Pó 1998a; Pó sfai et al., 1998b). The de novo synthesis of non-magnetic crystalline iron sulfide precursors to greigite aligned along the magnetosome chain indicates that chain formation within the cell does not involve magnetic interactions. Interestingly, under the strongly reducing, sulfidic conditions at neutral pH in which the greigite-producing magnetotactic bacteria are found (Bazylinski et al., 1990; Bazylinski and Frankel, 1992), greigite particles would be expected to transform into pyrite (Berner, 1967; Berner, 1970) which has not been unequivocally identified in magnetotactic bacteria. It is not known if and how cells prevent this transformation. As with magnetite, three particle morphologies of greigite have been observed in magnetotactic bacteria (Fig. 14): 1) cubo-octahedral (the equilibrium form of face-centered cubic greigite) (Heywood et al., 1990; Heywood et al., 1991); 2) pseudo-rectangular prismatic as shown in Fig. 14 and 12e–f (Heywood et al., 1990; Heywood et al.,

CHAPTER 1.26

100 nm

a b

200 nm

c

50 nm Fig. 14. Morphologies of intracellular greigite (Fe3S4) particles produced by magnetotactic bacteria. (a) Brightfield STEM image of cubo-octahedra in an unidentified rodshaped bacterium collected from the Neponset River estuary, Massachusetts, USA. (b) Brightfield STEM image of rectangular prismatic particles in an unidentified rod-shaped bacterium collected from the Neponset River estuary, Massachusetts, USA. (c) Brightfield TEM image of tooth-shaped and rectangular prismatic particles from the many-celled magnetotactic prokaryote (MMP), courtesy of M. Pó sfai and P. R. Buseck.

CHAPTER 1.26

Magnetotactic Bacteria

1991); and 3) tooth-shaped (Pó sfai et al., 1998a; Pó sfai et al., 1998b). Like that of their magnetite counterparts, the morphology of the greigite particles also appears to be species- and/or strain-specific, although confirmation of this observation will require controlled studies of pure cultures of greigite-producing magnetotactic bacteria, none of which is currently available. One clear exception to this rule is the MMP (Farina et al., 1983; Bazylinski et al., 1990; Bazylinski et al., 1993a; Mann et al., 1990b; Rodgers et al., 1990a; 1990b; Bazylinski and Frankel, 1992). This unusual microorganism, found in salt marsh pools all over the world and some deep sea sediments, has been found to contain pleomorphic, pseudorectangular prismatic, tooth-shaped, and cubo-octahedral greigite particles. Some of these particle morphologies are shown in Fig. 7 and 14c. Therefore the biomineralization process(es) appear(s) to be more complicated in this organism than in the rods with greigite-containing magnetosomes or in magnetite-producing, magnetotactic bacteria.

chain direction. Both particle morphologies have been found in organisms with single mineral component chains (Mann et al., 1987a; Mann et al., 1987b; Heywood et al., 1990; Heywood et al., 1991), which suggests that the magnetosome membranes surrounding the magnetite and greigite particles contain different nucleation templates and that there are differences in magnetosome vesicle biosynthesis. Thus, it seems likely that two separate sets of genes control the biomineralization of magnetite and greigite in this organism.

Phylogeny The phylogeny of many morphotypes of magnetotactic bacteria, including both those in pure culture and those collected from natural environments, has been determined by sequencing their 16S rRNA genes. To date, representatives of the magnetotactic prokaryotes are phylogenetically associated with three major lineages within the Bacteria. Although most are located within the Proteobacteria, “Magnetobacterium bavaricum” is affiliated with another phylum, the newly designated Nitrospira group. Those within the Proteobacteria are distributed among the delta- and alpha-subclasses. The uncultured greigite-producing, MMP and the magnetite-producing, sulfate-reducing magnetotactic bacterium RS-1, which is available in pure culture, are located in the delta-subclass, whereas members of the genus Magnetospirillum and various vibrios and coccoid magnetotactic bacteria, all of which produce magnetite, belong to the alpha-subclass (Fig. 15). Although these results suggest that the trait of magnetotaxis in bacteria has multiple evolutionary origins (DeLong et al., 1993), it is also possible that the ability of magnetosome

Magnetite and Greigite Crystals in a Single Bacterium One slow-swimming, rod-shaped bacterium, collected from the OATZ from the Pettaquamscutt Estuary, was found to contain arrowhead-shaped crystals of magnetite and rectangular prismatic crystals of greigite coorganized within the same chains of magnetosomes (this organism usually contains two parallel chains of magnetosomes) (Bazylinski et al., 1993b; Bazylinski et al., 1995). In cells of this uncultured organism, the magnetite and greigite crystals occur with different, mineralspecific morphologies and sizes and are positioned with their long axes oriented along the

Rhodopseudomonas palustris Paracoccus denitriticans Agrobacterium vitis

Brevundimonas diminuta Magnetospirillum magnetotacticum M. gryphiswaildense Sphingomonas capsulata Magnetotactic vibria MV1

Magnetotactic coccus MC1 mabrj58

Oceanospirillum pusillum

Fig. 15. Phylogenetic tree based on 16S rRNA sequences showing the positions of cultured and uncultured magnetotactic bacteria within the alpha-subclass of Proteobacteria. Sequences of uncultured magnetotactic bacteria retrieved from freshwater habitats are in blue, from marine habitats in red, and from a lagoon in pink.

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macpa11 mabrj12 macpa19 Uncultured macpa119 magnetotactic bacteria macca81

Rhodospirillum sallnarum Rhodopila giobiformis

macth12mabca92 macth24 macca13 macmp17 macca38

10%

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formation was spread among various phylogenetic groups of bacteria and even eukaryotes by lateral gene transfer. To date, most of the 16S rRNA sequences of magnetotactic bacteria retrieved from environmental samples form a deep-branching group within the alpha-subclass (Fig. 15). This phylogenetic assemblage consists (up to now) exclusively of bacteria displaying magnetotaxis. Similarity values of 16S rRNA sequences within this monophyletic group of magnetotactic bacteria range from 88.0 to 99.3%. Using in situ hybridization with fluorescently-labeled oligonucleotide probes, it was demonstrated that members of this coherent phylogenetic cluster represent the dominant fraction of magnetotactic bacteria in many environments like lagoons, marine and freshwater sediments (Spring et al., 1992; Spring et al., 1994; Spring et al., 1998). Magnetotactic bacterial morphotypes in this group, as evidenced by in situ hybridization, are mainly represented by coccoid to ovoid bacteria, but also include one rod- to vibrio-shaped bacterium (mabcs92; Fig. 15). Despite continuous effort in several laboratories, most members of this group have resisted attempts (in several laboratories) at isolation to axenic culture. One major reason may be their adaptation to and requirement for gradient systems not easily replicated in synthetic growth media. The only exception is the marine magnetotactic coccus strain MC-1, which can be cultivated in a synthetic oxygen gradient medium.

Cultivation and Physiology The Genus Magnetospirillum Taxonomy Magnetospirilla are found in freshwater habitats where they usually occur in low numbers as, for example, compared with the magnetotactic cocci. These clockwise spirilla have dimensions of 0.2–0.7 by 1–20 mm and display an axial magneto-aerotaxis, at least when grown in liquid culture. The genus Magnetospirillum currently comprises the two validly described species, M. magnetotacticum and M. gryphiswaldense, and several partially characterized strains. M. magnetotacticum was the first magnetotactic bacterium isolated and grown in pure culture and was originally assigned to the genus Aquaspirillum based on a number of phenotypic characteristics (Maratea and Blakemore, 1981). At that time, this genus contained a large number of phylogenetically diverse, nonphototrophic, freshwater spirilla with the type species A. serpens phylogenetically located among the beta-subclass of the Proteobacteria. Phylogenetic analyses of Magnetospirillum strains later revealed that they all belong to a phylogenetic

CHAPTER 1.26

branch within the alpha-subclass of the Proteobacteria and are closely related to phototrophic spirilla of the genus Phaeospirillum (Fig. 15; Burgess et al., 1993; Schü ler et al., 1999). Therefore, it was justified to propose the new genus Magnetospirillum for these strains (Schleifer et al., 1991). Members of this genus can be distinguished from other freshwater spirilla by their ability to produce membrane-enveloped cubooctahedral magnetite crystals, averaging about 42 nm in diameter (Balkwill et al., 1980; Schleifer et al., 1991), arranged in a single chain within the cell. Other characteristic traits of members of this genus include bipolar monotrichous flagellation and a preference for microoxic growth conditions. Several strains of magnetospirilla can grow also anaerobically with nitrate as terminal electron acceptor or aerobically with atmospheric concentrations of oxygen. Magnetite synthesis appears to only occur under microaerobic conditions in most species while Magnetospirillum strain AMB-1 appears to synthesize magnetite under anaerobic conditions as well (Matsunaga and Tsujimura, 1993). Preferred substrates are intermediates of the tricarboxylic acid cycle and acetate. Carbohydrates are not utilized. Catalase and oxidase may be present or not. The guanine-plus-cytosine content of DNA ranges from 64 to 71 mol% (Burgess et al., 1993). Biochemistry and Molecular Biology of Magnetosome Formation There has been much interest in the elucidation of magnetosome formation because the crystals synthesized by magnetotactic bacteria are of great structural perfection, have consistent particle morphologies and narrow size distributions, possible indications that the particles may have novel magnetic, physical and/or electrical properties. Understanding the factors controlling the biomineralization of iron in magnetosome synthesis within bacteria could also be helpful for the elucidation of similar processes in animals and man or for the artificial synthesis of biominerals. Despite the dedicated and elaborate efforts in studying magnetosome synthesis in bacteria, published results are rather sparse. This is partly due to the lack of a significant number of magnetotactic bacteria strains and the difficulty in culturing them reproducibly in the laboratory, which would be a prerequisite for the establishing of biochemical or genetic model systems. Nevertheless, some interesting results have been obtained using the few available, but fastidious Magnetospirillum strains. In general, the bacterial magnetite synthesis can be divided into three steps. Initially, extracellular iron has to be transported across the cell wall to the inside of the cell. Once within the cell, iron must accumulate in specialized compart-

CHAPTER 1.26

ments, the magnetosome vesicles. There, the iron presumably precipitates and transforms or grows into a single-magnetic-domain magnetite crystal with a specific morphology. It is assumed that the membrane vesicle is synthesized prior to the precipitation of iron but since there is currently little evidence to support this idea, it is possible that the precipitation of iron and crystal nucleation occurs first and the magnetosome membrane then forms around the growing crystal. The uptake of iron from the surrounding environment by cells of Magnetospirillum strains has been analyzed by several groups (Paoletti and Blakemore, 1986; Nakamura et al., 1993b; Schü ler and Baeuerlein, 1996; Schü ler and Baeuerlein, 1998). Generally, the results suggest that iron is taken up by the cell in the ferric form and transported across the membrane by an energy-dependent reductive process. Ironbinding siderophores were thought to be involved in iron uptake by M. magnetotacticum (Paoletti and Blakemore, 1986), which appeared to produce a hydroxamate siderophore under high, but not low, iron conditions. However, this finding was never confirmed by other laboratories. Spent culture fluid stimulates the uptake of ferric iron in M. gryphiswaldense although there was no evidence for the production of a siderophore by this species. This stimulation may be due to the production of unknown compounds, produced by cells during growth, which mediate iron uptake by an unrecognized novel mechanism (Schü ler and Baeuerlein, 1996). In this respect, it is noteworthy that most magnetotactic bacteria are adapted to microenvironments, like the oxic-anoxic transition zone of sediments, where soluble iron is available to the cell in sufficient quantities for magnetite synthesis (generally about 10–20 mM iron; Blakemore et al., 1979). Thus, magnetotactic bacteria probably have no need for high-affinity transport systems like many other aerobic bacteria growing under iron deficient conditions. This is consistent with experiments performed with cells of M. gryphiswaldense. Under iron deficient conditions, cells of M. gryphiswaldense do not or cannot distinguish between the use of incorporated iron either as a cofactor for cellular proteins or for magnetite synthesis and store this essential element as an inorganic mineral, magnetite, at the expense of their own growth (Schü ler and Baeuerlein, 1996). The marine magnetotactic vibrio, strain MV-1, behaves similarly. The fate of iron taken up by cells was studied by Frankel et al. (1983) in M. magnetotacticum using Fe57 Moessbauer spectroscopy. It was proposed that the ferrous iron taken up by cells is immediately reoxidized to form a low-density hydrous Fe(III) oxide. It is not yet clear if this step takes place in the cytoplasm or in the mag-

Magnetotactic Bacteria

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netosome vesicles. How iron is transported from the cell membrane into the magnetosome vesicle is also not known. Iron is precipitated within the magnetosome vesicle presumably through a dehydration step as ferrihydrite (a high-density Fe(III) hydroxide). Finally, magnetite (Fe3O4) is produced by the reduction of one-third of the Fe(III) ions in ferrihydrite and further dehydration steps. The crystallization process(es) involved in magnetite formation is apparently linked closely to the magnetosome membrane and may be controlled by specific proteins present in this membrane. The chemical transformation of amorphous Fe(III) precursors to crystalline magnetite is sensitive to environmental conditions like ion concentration, pH and redox potential (Mann et al., 1990c) which have to be therefore precisely regulated by the magnetosome membrane or by conditions within the magnetosome membrane vesicle. Growth of the magnetite crystal, i.e., its orientation, shape and size, must also be under strict control because these characteristics are specific for one strain and/or species of bacteria and to a great extent independent from the growth conditions (Bazylinski et al., 1994). Because the magnetosome membrane seems to play a key role in the synthesis of magnetite crystals, its structure and composition has been analyzed in several studies. By analyzing the magnetosome membrane in this way, some clues relating to how magnetite biomineralization occurs within the cell may be found. Gorby et al. (1988) showed that the magnetosome membrane in M. magnetotacticum has an architecture similar to that of the cytoplasmic membrane and consists of a lipid bilayer and numerous proteins, some of which appear to be unique to the magnetosome membrane. Okuda et al. (1996) found three proteins with molecular weights of 12, 22 and 28 kDa, specifically associated with the magnetosome membrane in M. magnetotacticum. They successfully identified and sequenced the gene encoding for the 22 kDa protein, which was found to belong to a family of protein import receptors common in mitochondria and peroxisomes. The role of this protein in magnetosome synthesis remains unclear however. A gene likely involved in magnetite synthesis was identified and characterized by Matsunaga et al. (1992). They used a genetic approach using the microorganism Magnetospirillum strain AMB-1, which forms colonies of magnetite-forming cells on agar surfaces, thereby facilitating the screening for nonmagnetic mutants. The gene, designated magA, encodes for a membrane protein showing sequence similarities to some cation efflux proteins. Based on experiments with the recombinant protein, it was proposed that the MagA

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protein plays a role in the energy-dependent transport of iron across membranes. Enrichment and Isolation Magnetotactic spirilla have been repeatedly isolated from various freshwater habitats, so that it is possible to give some guidelines for their succesful enrichment and isolation. Several morphotypes of magnetotactic bacteria can be enriched in the laboratory by putting mud and overlying water from a sampling site into aquaria or jars, which are loosely covered and stored in dim light. After several days to weeks, the number of magnetotactic bacterial cells generally increases significantly. Magnetotactic spirilla, however, are in most cases not among the dominating morphotypes and therefore are only rarely detected using light microscopy. Consequently, the usefulness of this method for the enrichment of representatives of the genus Magnetospirillum remains questionable. To date, no selective growth media are known for the cultivation of magnetotactic spirilla, so that a successful isolation procedure will in most cases depend on the purity of the inoculum. Because of the magnetic dipole moment of these bacteria, physical separation from nonmagnetotactic contaminants is possible. A commonly used method for the separation of magnetotactic bacteria from sediment samples was described by Moench and Konetzka (1978). They concentrated magnetotactic bacterial cells using a bar magnet (e.g., stirring bar) fixed to the outer wall of a jar filled with sediment and water. Directed magnetotactic bacteria eventually accumulate at the side of the jar and become concentrated enough to form (opposite to the magnet) a brownish spot from where they can be transferred into a sterile cap using a Pasteur pipette. The sample containing the concentrated magnetotactic cells still cotains non-magnetotactic bacteria, so that a further purification step is advisable before it is used as an inoculum for growth media. The “capillary racetrack” devised by Spormann and Wolfe (Wolfe et al., 1987) has been successfully used for this purpose (Schü ler et al., 1999). All isolated strains of magnetospirilla, with the possible exception of Magnetospirillum strain AMB-1, appear to prefer low oxygen tensions for growth and magnetite synthesis. Thus the creation and maintenance of microoxic conditions in growth media is especially important for the isolation of these organisms starting from small inocula. The growth medium should contain 10– 20% of sterilized mud or water from the respective habitat and low concentrations of agar to allow the establishment of a semisolid oxygen gradient. Suitable carbon sources are intermediates of the tricarboxylic acid cycle, e.g., malate or

CHAPTER 1.26

succinate. An oxygen-sulfide gradient medium was successfully used by Schü ler et al. (1999) for the effective isolation of magnetotactic spirilla from a freshwater pond. Screw-capped culture tubes are filled with 1 ml of solid sulfide agar (4 mM Na2S, 1.5% agar, pH 7.4) and overlaid with 10 ml of slush-agar. The slush-agar consists of (per 800 ml deionized water): 200 ml of filtered pond water, 1 ml of vitamin elixir, 2 ml of mineral elixir (Wolin et al., 1963), 0.05 g, sodium succinate, 0.05 g, yeast extract, 0.05 g, NH4Cl, 0.05 g, MgSO4, 0.5 mM potassium phosphate buffer (pH 7.0); 2 mg of resazurin and 2 g of agar. After adjusting the pH to 7.0 and autoclaving, sterile solutions of ferric citrate and neutralized cysteine·HCl are added (final concentrations 10 mM and 0.01%, respectively). The culture tubes can be inoculated after the establishment of sulfide and oxygen gradients within the medium, which takes about 24 hours. Several days to weeks of incubation at room temperature in the dark may be required until growth becomes apparent, usually as fluffy pinpoint colonies. Although the original inoculum always contains various types of magnetotactic bacteria, in most cases only magnetotactic spirilla grow and are eventually isolated in pure culture. Modifications of this medium may eventually prove useful for the isolation of hitherto uncultured types of magnetotactic bacteria. Following isolation, most strains of magnetospirilla can be cultured in liquid media without added water or mud from the sampling site. However, the gas composition of the headspace of the cultures is crucial for good growth and magnetite synthesis by most species. The maximum concentration of oxygen allowing growth and/or magnetite synthesis differs among the described Magnetospirillum strains. M. magnetotacticum grows optimally and produces the highest number of magnetosomes at an oxygen tension of 1% in the headspace and tolerates higher initial oxygen concentrations only if a large number of cells is inoculated into the growth medium. In contrast, Magnetospirillum strain AMB-1 grows (but does not produce magnetosomes) aerobically under atmospheric concentrations of oxygen (approximately 21% O2; Matsunaga et al., 1991b). Magnetite synthesis is inhibited by all Magnetospirillum strains when cells are cultured under oxygen concentrations above 2 to 6% (Blakemore et al., 1985; Schü ler and Baeuerlein, 1998).

Strain MV-1, a Facultatively Anaerobic Magnetotactic Vibrio A marine magnetotactic vibrioid to helicoid bacterium, strain MV-1, was isolated by Bazylinski et al. (1988). Cells of strain MV-1 are small, rang-

CHAPTER 1.26

Magnetotactic Bacteria

a

1 µm

b

100 nm Fig. 16. Brightfield TEM image of negatively stained cell and magnetosomes of strain MV-1. (a) Cell stained with uranyl acetate showing a single polar flagellum and a chain of magnetite-containing magnetosomes. (b) Preparation of purified magnetosomes from strain MV-1 stained with 2% aqueous sodium phosphotungstate, pH 7.0. The “magnetosome membrane” is visualized as an electron-lucent area surrounding each individual crystal and is easily removed with detergents such as sodium deodecyl sulfate.

ing from 1–5 mm by 0.2–0.5 mm, and possess a single, unsheathed, polar flagellum (Fig. 16a). Cells grow and synthesize pseudohexahedral prismatic crystals of magnetite, averaging 53 by 35 nm in size (Fig. 16b; Sparks et al., 1990), in their magnetosomes microaerobically and anaerobically, with nitrous oxide as the terminal electron acceptor. Cells appear to produce more magnetite under anaerobic conditions than under microaerobic conditions (Bazylinski et al., 1988) and, like M. magnetotacticum, synthesize a number of magnetosome membrane proteins that are not present in other cellular fractions (Dubbels et al., 1998). A stable, spontaneous nonmagnetotactic mutant strain of MV-1 that does not produce magnetosomes has recently been isolated and partially characterized (Dubbels and Bazylinski, 1998). Strain MV-1 is nutritionally versatile being able to grow chemoorganoheterotrophically with organic and some amino acids as carbon and

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energy sources, and chemolithoautotrophically with thiosulfate or sulfide as energy sources oxidizing them to sulfate, and carbon dioxide as the sole carbon source (Kimble and Bazylinski, 1996). Cells produce intracellular sulfur deposits when grown with sulfide (Kimble and Bazylinski, 1996). As do virtually all aerobic chemolithoautotrophic bacteria, strain MV-1 uses the CalvinBenson cycle for autotrophic carbon dioxide fixation (McFadden and Shively, 1991). Cell-free extracts from thiosulfate-grown cells of strain MV-1 show ribulose bisphosphate carboxylase/ oxygenase (rubisCO) activity (Kimble and Bazylinski, 1996), and recently (Dean and Bazylinski, 1999a) the gene for a form II rubisCO enzyme (cbbM) was cloned and sequenced from strain MV-1. There was no evidence for a cbbL gene (encodes for form I rubisCO enzymes) in DNA hybridization analyses despite using cbbL gene probes from several different organisms. Because many uncultured magnetotactic bacteria collected from natural habitats thrive in oxygen-sulfide inverse gradients, as previously mentioned, and contain internal sulfur deposits (Moench, 1988; Spring et al., 1993; Frankel and Bazylinski, 1994; Iida and Akai, 1996; Kimble and Bazylinski, 1996), it seems many species are likely chemolithoautotrophs that obtain energy from the oxidation of sulfide and perhaps other reduced sulfur sompounds. Using pulsed-field gel electrophoresis (PFGE), the genome of strain MV-1 was found to consist of a single, circular chromosome of approximately 3.7 Mb (Dean and Bazylinski, 1999b). There was no evidence of linear chromosomes or extrachromosomal DNA such as plasmids. The guanine-pluscytosine content of the DNA of this strain is 52.9 mol% as determined by HPLC and 53.5 mol% by Tm. A virtually identical strain to strain MV-1, designated MV-2, was isolated from the Pettaquamscutt Estuary (DeLong et al., 1993; Meldrum et al., 1993b). Cells of this strain produce the same morphological type of magnetite crystals as strain MV-1 (Meldrum et al., 1993b) and display many of the same phenotypic traits as strain MV-1 (such as anaerobic growth with nitrous oxide as a terminal electron acceptor, heterotrophic growth with organic and amino acids, and chemolithoautotrophic growth on reduced sulfur compounds). However, strain MV-2 shows slightly different restriction fragment patterns in pulsed-field gels than strain MV-1 using the same restriction enzymes (Dean and Bazylinski, 1999b). As with strain MV-1, the genome of strain MV-2 consists of a single, circular chromosome of a similar size, about 3.6 Mb (Dean and Bazylinski, 1999b). The guanine-plus-cytosine content of the DNA of this strain is 56.2 mol% as determined by HPLC and 56.6% by Tm.

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Strain RS-1, a Sulfate-Reducing Magnetotactic Bacterium It was thought for a long time that all magnetotactic bacteria are obligate or facultative microaerophiles (Magnetospirillum strain AMB1 and the marine vibrio, strain MV-1, grow anaerobically with nitrate and nitrous oxide, respectively, as well as with oxygen) adapted to the microoxic zone of their environment. With the isolation of an obligately anaerobic strain from a sulfidic freshwater habitat by Sakaguchi et al. (1993), this assumption is clearly incorrect. Cells of this organism, designated strain RS-1, are 0.9–1.5 by 3–5 mm with a helicoid to rodshaped morphology and possess a single polar flagellum. They exhibit an axial magnetotaxis coupled with a strong anaerotaxis reflecting their obligate anaerobic metabolism. According to the revised model of magnetotaxis, they may have developed an axial magnetotaxis because they do not have to oscillate between microoxic and anoxic zones of their habitat, which would select for polar magnetotaxis. Strain RS-1 is a dissimilatory sulfate-reducing, chemoorganoheterotrophic bacterium that utilizes a variety of organic substrates, e.g., pyruvate, lactate, ethanol and fumarate). Cells can use sulfate or fumarate as electron acceptor but not oxygen. They are catalase positive and oxidase negative. The guanine-plus-cytosine content of DNA was determined by HPLC to be 66 mol%. Sequencing of the 16S rRNA gene of strain RS-1 showed that it is phylogenetically affiliated to the delta-subclass of the Proteobacteria (Kawaguchi et al., 1995). The nearest neighbors in a phylogenetic tree are members of the genus Desulfovibrio, typical representatives of the obligately anaerobic, dissimilatory sulfate-reducing bacteria. In contrast to Desulfovibrio sp., cells of strain RS-1 are able to produce intracellular bean-shaped crystals of magnetite, responsible for its magnetotactic response. Consequently, the production of magnetosomes consisting of magnetite is found in bacteria belonging to three different phylogenetic groups, viz. the a- and dsubclasses of Proteobacteria and the Nitrospira group (“Magnetobacterium bavaricum”), indicating multiple evolutionary origins of intracellular magnetite synthesis or lateral gene transfer between different phylogenetic groups.

Other Magnetotactic Strains in Pure Culture Several other pure cultures of magnetotactic bacteria exist, but they appear to be obligate microaerophiles and grow poorly (D. A. Bazylinski, unpublished results). Hence, very little is known about them. Strain MC-1 (Fig. 17), a

CHAPTER 1.26

0.5 µm

fb

p

s

m

Fig. 17. Brightfield TEM image of a cell of the bilophotrichously flagellated marine coccus strain MC-1 negatively stained with uranyl acetate. Note the two flagellar bundles (fb), the presence of pili (p), sulfur globules(s), and chain of Fe3O4-containing magnetosomes (m).

marine biliophotrichous coccus, was isolated from water collected from the Pettaquamscutt Estuary, a chemically-stratified semi-anaerobic basin in Rhode Island, USA. Cells of this strain produce pseudohexahedral prisms of magnetite, averaging 72 by 70 nm in size (when grown autotrophically), and grow chemolithoautotrophically with thiosulfate or sulfide as an electron and energy source (Meldrum et al., 1993a; Frankel et al., 1997). Cells may also be able to grow chemoorganoheterotrophically. Like all magnetotactic cocci observed, cells of strain MC-1 show polar magneto-aerotaxis regardless of whether they are grown in liquid or semi-solid oxygen gradient media. This strain has a genome size of approximately 4.5 Mb as determined by pulsed-field gel electrophoresis (Dean and Bazylinski, 1999b). The guanine-pluscytosine content of the DNA of strain MC-1, as determined by HPLC, is 55.8 mol%. This organism has not been completely characterized and described. Strain MV-4 (Fig. 18), a small marine spirillum, was isolated from sulfide-rich mud and water collected from School Street Marsh, Woods Hole, Massachusetts, USA. Cells of this strain produce elongated octahedrons of magnetite, averaging 61 by 52 nm in size, and grow chemolithoautotrophically with thiosulfate or chemoorganoheterotrophically with succinate (Meldrum et al., 1993b). Unlike most freshwater magnetotactic spirilla, this strain shows polar magneto-aerotaxis at least when grown in semisolid oxygen gradient media. Like strain MC-1,

CHAPTER 1.26

fb

Magnetotactic Bacteria

p

0.5 µm

Fig. 18. Brightfield TEM image of a cell of the marine spirillum strain MV-4 negatively stained with uranyl acetate showing bipolar flagellation and a chain of Fe3O4-containing magnetosomes.

this strain has not been completely characterized and described.

Biotechnological Applications It was not long after the discovery of magnetotactic bacteria that publications of physical studies and of commercial and medical applications involving the magnetotactic cells, isolated magnetosomes and/or magnetite crystals began to appear. It is clear that magnetotactic bacterial cells and their magnetic crystals have novel physical, magnetic and possibly electrical properties. In addition, in certain types of applications, bacterial magnetite offers several advantages compared to chemically synthesized magnetite. Bacterial magnetosome particles, unlike those produced chemically, have a consistent shape, a narrow size distribution within the single magnetic domain range, and a membrane coating consisting of lipids and proteins. The magnetosome envelope allows for easy couplings of bioactive substances to its surface, a characteristic important for many applications. Magnetotactic bacterial cells have been used to determine south magnetic poles in meteorites and rocks containing fine-grained magnetic minerals (Funaki et al., 1989; Funaki et al., 1992) and for the separation of cells after the introduction of magnetotactic bacterial cells into granulocytes and monocytes by phagocytosis (Matsunaga et al., 1989). Magnetotactic bacterial magnetite crystals have been used in studies of magnetic domain analysis (Futschik et al., 1989) and in many commercial applications including: the immobilization of enzymes (Matsunaga and Kamiya, 1987); the formation of magnetic antibodies in various fluoroimmunoassays

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(Matsunaga et al., 1990) involving the detection of allergens (Nakamura and Matsunaga, 1993a) and squamous cell carcinoma cells (Matsunaga, 1991a), and the quantification of IgG (Nakamura et al., 1991); the detection and removal of Escherichia coli cells with a fluorescein isothiocyanate conjugated monoclonal antibody, immobilized on magnetotactic bacterial magnetite particles (Nakamura et al., 1993c); and the introduction of genes into cells, a technology in which magnetosomes are coated with DNA and “shot” using a particle gun into cells that are difficult to transform using more standard methods (Matsunaga, 1991a). Unfortunately, the prerequisite for any large scale commercial application is mass cultivation of magnetotactic bacteria or the introduction and expression of the genes responsible for magnetosome synthesis into a bacterium, e.g., E. coli, that can be grown relatively cheaply to extremely large yields. Although some progress has been made, the former has not been achieved with the available pure cultures. Acknowledgements. We thank F.C. Meldrum, M. Pó sfai, P.R. Buseck, and M. Hanzlik for use of figures and L. Cox, D. Schü ler and R.B. Frankel for helpful discussions and suggestions concerning this chapter. S.S. is grateful to K.-H. Schleifer and the Deutsche Forschungsgemeinschaft (DFG) for continuous support. Research in the laboratory of D.A.B. is supported by U.S. National Science Foundation grant CHE9714101 and U.S. National Aeronautics and Space Administration grant NAG 9-1115.

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Bazylinski, D. A., B. R. Heywood, S. Mann, and R. B. Frankel. 1993b. Fe3O4 and Fe3S4 in a bacterium. Nature 366:218–219. Bazylinski, D. A., A. Garratt-Reed, and R. B. Frankel. 1994. Electron-microscopic studies of magnetosomes in magnetotactic bacteria. Microscopy Res. Tech. 27:389– 401. Bazylinski, D. A., R. B. Frankel, B. R. Heywood, S. Mann, J. W. King, P. L. Donaghay, and A. K. Hanson. 1995. Controlled biomineralization of magnetite (Fe3O4) and greigite (Fe3S4) in a magnetotactic bacterium. Appl. Environ. Microbiol. 61:3232–3239. Berner, R. A. 1967. Thermodynamic stability of sedimentary iron sulfides. Am. J. Sci. 265:773–785. Berner, R. A. 1970. Sedimentary pyrite formation. Am. J. Sci. 268:1–23. Berner, R. A. 1974. Iron sulfides in Pleistocene deep Black Sea sediments and their palaeooceanographic significance. In: E. T. Degens, and D. A. Ross (Eds.) The Black Sea: Geology, Chemistry and Biology. AAPG Memoirs 20:American Association of Petroleum Geologists. Tulsa, OK. 524–531. Bertani, L. E., J. S. Huang, B. A. Weir, and J. L. Kirschvink. 1997. Evidence for two types of subunits in the bacterioferretin of Magnetospirillum magnetotacticum. Gene 201:31–36. Blakemore, R. P. 1975. Magnetotactic bacteria. Science 190:377–379. Blakemore, R. P., D. Maratea, and R. S. Wolfe. 1979. Isolation and pure culture of a freshwater magnetic spirillum in chemically defined medium. J. Bacteriol. 140:720–729. Blakemore, R. P. 1982. Magnetotactic bacteria. Ann. Rev. Microbiol. 36:217–238. Blakemore, R. P., K. A. Short, D. A. Bazylinski, C. Rosenblatt, and R. B. Frankel. 1985. Microaerobic conditions are required for magnetite formation within Aquaspirillum magnetotacticum. Geomicrobiol. J. 4:53– 71. Blakemore, R. P., N. A. Blakemore, D. A. Bazylinski, and T. T. Moench. 1989. Magnetotactic bacteria. In: J. T. Staley et al. (Eds.) Bergey’s Manual of Systematic Bacteriology. 3:Williams and Wilkins. Baltimore, MD. 1882–1889. Bulte, J. W. M., and R. A. Brooks. 1997. Magnetic nanoparticles as contrast agents for imaging. In: U. Häfeli, W. Schü tt, J. Teller, and M. Zborowski (Eds.) Scientific and Clinical Applications of Magnetic Carriers. Plenum Press. New York, NY. 527–543. 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 rRNA sequences. J. Bacteriol. 175:6689–6694. Bulte, J. W. M., and R. A. Brooks. 1997. Magnetic nanoparticles as contrast agents for imaging. Häfeli, U., Schü tt, W., Teller, J., Zborowski, M.Scientific and clinical applications of magnetic carriers. Plenum Press. New York, 527–543. Butler, R. F., and S. K. Banerjee. 1975. Theoretical singledomain grain size range in magnetite and titanomagnetite. J. Geophys. Res. 80:4049–4058. Chang, S.-B. R., and J. L. Kirschvink. 1989a. Magnetofossils, the magnetization of sediments, and the evolution of magnetite biomineralization. Ann. Rev. Earth Planet Sci. 17:169–195.

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CHAPTER 1.26 Spormann, A. M., and R. S. Wolfe. 1984. Chemotactic, magnetotactic, and tactile behaviour in a magnetic spirillum. FEMS Microbiol. Lett. 22:171–177. Spring, S., R. Amann, W. Ludwig, K. H. Schleifer, and N. Petersen. 1992. Phylogenetic diversity and identification of nonculturable magnetotactic bacteria. Syst. Appl. Microbiol. 15:116–122. Spring, S., R. Amann, W. Ludwig, K. H. Schleifer, H. van Gemerden, and N. Petersen. 1993. Dominating role of an unusual magnetotactic bacterium in the microaerobic zone of a freshwater sediment. Appl. Environ. Microbiol. 59:2397–2403. Spring, S., R. Amann, W. Ludwig, K. H. Schleifer, D. Schü ler, K. Poralla, and N. Petersen. 1994. Phylogenetic analysis of uncultured magnetotactic bacteria from the alphasubclass of Proteobacteria. Syst. Appl. Microbiol. 17:501–508. Spring, S., U. Lins, R. Amann, K. H. Schleifer, L. C. S. Ferreira, D. M. S. Esquivel, and M. Farina. 1998. Phylogenetic affiliation and ultrastructure of uncultured magnetic bacteria with unusually large magnetosomes. Arch. Microbiol. 169:136–147. Steinberger, B., N. Petersen, H. Petermann, and D. G. Weiss. 1994. Movement of magnetic bacteria in time-varying magnetic fields. J. Fluid Mech. 273:189–211. Stolz, J. F., S.-B. R. Chang, and J. L. Kirschvink. 1986. Magnetotactic bacteria and single-domain magnetite in hemipelagic sediments. Nature 321:849–851. Stolz, J. F., D. R. Lovley, and S. E. Haggerty. 1990. Biogenic magnetite and the magnetization of sediments. J. Geophys. Res. 95:4355–4361. Stolz, J. F. 1993. Magnetosomes. J. Gen. Microbiol. 139:1663– 1670. Thornhill, R. H., J. G. Burgess, T. Sakaguchi, and T. Matsunaga. 1994. A morphological classification of bacteria containing bullet-shaped magnetic particles. FEMS Microbiol. Lett. 115:169–176. Towe, K. M., and T. T. Moench. 1981. Electron-optical characterization of bacterial magnetite. Earth Planet. Sci. Lett. 52:213–220. Vali, H., O. Fö rster, G. Amarantidis, and N. Petersen. 1987. Magnetotactic bacteria and their magnetofossils in sediments. Earth Planet. Sci. Lett. 86:389–426. Wolfe, R. S., R. K. Thauer, and N. Pfennig. 1987. A capillary racetrack method for isolation of magnetotactic bacteria. FEMS Microbiol. Lett. 45:31–35. Wolin, E. A., M. J. Wolin, and R. S. Wolfe. 1963. Formation of methane by bacterial extracts. J. Biol. Chem. 238:2882–2886.

Prokaryotes (2006) 2:863–892 DOI: 10.1007/0-387-30742-7_27

CHAPTER 1.27 suon imuL

a i re t caB

Luminous Bacteria PAUL V. DUNLAP AND KUMIKO KITA-TSUKAMOTO

Introduction and Historical Perspective “The smallest lamps in the world, luminous bacteria, are no different from ordinary bacteria except in their ability to luminesce.” —E. N. Harvey, 1940

The luminous bacteria are those bacteria that contain naturally acquired genes for light production, the lux genes. Most currently known luminous bacteria express the lux genes at high levels in laboratory culture (Fig. 1) or in nature, leading to the emission of easily visible levels of light. Bacterial light production is one of several biochemically distinct types of bioluminescence (Hastings, 1995). The existence of bacterial luminescence and of many of the luminous bacteria themselves has been known for some time. During the 1700s and 1800s, various animal products (such as meats, fish and eggs), the decaying bodies of marine and terrestrial animals, and even human wounds and corpses, were reported to produce light (Harvey, 1940; Harvey, 1952). Many years before those observations and long before bacteria were known to exist, Boyle (1668) demonstrated that the “uncertain shining of Fish,” the light coming from decaying fish, required air. Indeed, encounters with luminous objects and substances extend back to the beginnings of recorded history in Greece and China (Harvey, 1957), and they continue in modern times to be causes of concern and wonder. Many of these encounters can be attributed to the saprophytic or pathogenic growth of luminous bacteria on marine and terrestrial animals. According to Harvey (1940), J. F. Heller in 1854 was the first to give a name, Sarcina noctiluca, to the suspected responsible organism. As the science of bacteriology developed during the period from 1860 through 1910, individual types of light-producing bacteria were grown and distiguished from other bacteria, notably through the work of Bernhard Fischer (Fischer, 1887) and Martin Beijerinck (Beijerinck, 1889), among many others (Zobell, 1946; Harvey, 1952; Harvey, 1957). During the first half

of the 20th century, luminous bacteria were isolated from various habitats, the chemistry of bacterial light production, and culture requirements for growth and luminescence were characterized, and they were placed in the evolving system of microbial taxonomy (e.g., Zobell and Upham, 1944; Farghaly, 1950; Johnson, 1951). In the latter half of the 20th century, those efforts paralleled the growth of microbiology, incorporating the tools and knowledge developing from advances in biochemistry, physiology and genetics (Baumann and Baumann, 1977; Baumann and Baumann, 1981; Farmer and Hickman-Farmer, 1992; Hastings and Nealson, 1977; Hastings and Nealson, 1981; Hendrie et al., 1970; Nealson and Hastings, 1992; Singleton and Skerman, 1973). Much has been learned during the past 50 years about the enzymes and genes involved in bacterial light production and about the phylogeny and ecology of light-emitting bacteria. However, much remains to be learned about these topics and about the evolutionary origins and cellular functions of bacterial luminescence. In the past, the luminous bacteria were often considered to be a separate microbial group, distinguished by their distinctive and unifying phenotype, the production of light. They are seen now more properly as representative prokaryotes with much to reveal about the fundamental biology of bacteria. This view develops from (and is supported by) a deepening understanding of phylogenetic relationships and the realization that these bacteria are metabolically similar to other well-established bacteria (Baumann and Baumann, 1981; Baumann and Schubert, 1984a; Baumann et al., 1984b). Indeed, light-emission is a biochemical trait shared by several but not all species and strains of the genera Vibrio, Photobacterium, Shewanella and Photorhabdus. As members of these genera, the luminous bacteria are for the most part typical Gram negative bacteria similar in fundamental ways to terrestrial enterobacteria (Baumann and Baumann, 1977). They occur together with closely related nonluminous types in many habitats, responding in the same metabolic and physiological ways as other bacteria, and carrying out with them

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multitude of scientific advances and research opportunities opened up by the revelations of quorum-sensing control of luminescence suggest that studying luminous bacteria (as representative Gram-negative prokaryotes) and luminescence (as an integral aspect of their biology) will continue to reveal insights and themes of biological importance.

Biochemistry of Bacterial Light Production

Fig. 1. Bacterial luminescence. Colonies of the luminous marine bacterium V. fischeri growing on a seawater-based complete medium photographed by the light they produce. From Meighen and Dunlap (1993).

ecologically important activities unrelated to luminescence. Supporting this view is a growing appreciation that light production in luminous bacteria is tightly integrated with cellular metabolism and global gene regulation (Ulitzur and Dunlap, 1995; Callahan and Dunlap, 2000). Light production is sensitive to the physiological state of the cell, and expression of the lux genes, along with many other sets of genes of diverse functions, is coordinately regulated in response to that state. Therefore, despite its phenotypic distinctiveness, luminescence is not an independent biochemical activity of the cell; it is instead an integral feature of the biology of these bacteria. Studies of luminescence therefore are likely to reveal basic processes in Gram-negative prokaryotes. An example of how bacterial luminescence can lead to insights of fundamental importance in microbiology is quorum sensing. Previously called “autoinduction” and studied as the special cell density-dependent mechanism by which luminous bacteria control light production, quorum sensing has now been identified in many nonluminous Gram-negative bacteria, including several pathogens of animals and plants (Fuqua et al., 1996; Greenberg, 1997; Dunlap, 1997; Swift et al., 1999; Hastings and Greenberg, 1999). Early studies of luminous bacteria would not have led to predictions that autoinduction of luminescence would become a new bacterial regulatory paradigm (Nealson, 1999). However, the

Light emission in bacteria is catalyzed by luciferase, a heterodimeric protein of approximately 80 kD, composed of a (40-kDa) and b (37-kDa) subunits. Bacterial luciferase mediates the oxidation of reduced flavin mononucleotide (FMNH2) and a long-chain aliphatic aldehyde (RCHO) by molecular oxygen (O2) to produce blue-green light (Fig. 1) according to the following reaction. luciferase

FMNH 2 + O 2 + RCHO æ ææ æÆ FMN + H 2O + RCOOH + hv(490 nm) In the luminescence reaction, binding of FMNH2 by the enzyme is followed by interaction with O2 to form a luciferase-bound 4a-peroxyflavin. Association of this complex with aldehyde forms a highly stable intermediate, the slow decay of which results in oxidation of the FMNH2 and aldehyde substrates and the emission of light. Quantum yield for the reaction has been estimated at 0.1 to 1.0 photons. The reaction is highly specific for FMNH2, and the aldehyde substrate in vivo is likely to be tetradecanal. Synthesis of the long-chain aldehyde is catalyzed by a fattyacid reductase complex composed of three polypeptides, an NADPH-dependent acyl protein reductase (called “r,” 54 kDa), an acyl transferase (“t,” 33 kDa), and an ATP-dependent synthetase (“s,” 42 kDa). The complex has a stoichiometry of r4s4t2-4, and its activity is essential for the production of light in the absence of exogenously added aldehyde. The genes luxA and luxB for the a and b luciferase subunits and luxC, luxD and luxE for the r, s and t polypeptides of the fatty-acid reductase, respectively, are contiguous and coordinately expressed in all luminous bacteria examined to date. Furthermore, as described in a later section, luciferases from different species of luminous bacteria exhibit substantial sequence identity, consistent with a common evolutionary origin. For references and detailed information on the biochemistry of bacterial light production, the reader is directed to reviews by Hastings (1995), Hastings et al. (1985), Meighen (Meighen, 1988; Meighen, 1991) and Meighen and Dunlap (1993).

CHAPTER 1.27

Species and Phylogeny of Luminous Bacteria The currently known luminous bacteria are members of the genera Vibrio, Photobacterium, Shewanella and Photorhabdus (Table 1). These bacteria are Gram-negative g-Proteobacteria, nonsporulating, chemoorganotrophic heterotrophs, most of which are facultatively aerobic. Two of the marine luminous bacteria, however, Shewanella hanedai (Jensen et al., 1980) and Shewanella woodyi (Makemson et al., 1997), differ from the other luminous bacteria in using only a respiratory mode of metabolism. Detailed information on the metabolism, physiology and morphology of these bacterial groups and individual species can be found in Baumann and Baumann (1981), Baumann et al. (1984b), Farmer and Hickman-Brenner (1992), Boemare et al. (1993) and Forst et al. (1997). The luminous Photobacterium and Shewanella species and most of the luminous Vibrio species occur in the marine environment, whereas Photorhabdus species are terrestrial. Vibrio cholerae may be the only species with luminous strains occurring in brackish environments and freshwater. Certain of the species listed in Table 1 were described in the late 1990s (Makemson et al., 1997; Fischer-Le Saux et al., 1999). Furthermore, luminous strains of species previously described as nonluminous are being found. Examples include Vibrio salmonicida, a pathogen of salmonid fish (Fidopastis et al., 1999) and intensely luminous strains of Photobacterium angustum isolated from the Sea of Cortez (K. Kita-Tsukamoto et al., manuscript in preparation). Indeed, light production does not define a phylogenetically exclusive or consistent grouping. The genera Vibrio, Photobacterium and Shewanella contain many nonluminous species. Even species characterized as luminous can contain strains that do not produce light and that lack the genes necessary for light production. An example is a strain of Photorhabdus luminescens symbiotic with entomopathogenic nematodes (Akhurst and Boemare, 1986; Forst and Nealson, 1996). Adding to this complexity, some species or strains carry the lux genes and produce a high level of light under natural conditions but produce little or no light when grown in laboratory culture. Examples include luminous bacteria infecting crustaceans (Giard and Billet, 1889b) and strains of V. fischeri symbiotic with the Hawaiian sepiolid squid, Euprymna scolopes (Boettcher and Ruby, 1990). Furthermore, many nonluminous strains of V. cholerae carry lux genes (Palmer and Colwell, 1991; Ramaiah et al., 2000). Relevant to the question of which species

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and strains of bacteria produce light is the observation that luminescence often is not phenotypically stable. Strains luminous on primary isolation often become dim or dark in laboratory culture (Nealson and Hastings, 1979b; Akhurst, 1980; Silverman et al., 1989; Nealson and Hastings, 1992). Therefore, it is reasonable to assume that luminescence has been overlooked in many species, especially those represented primarily by laboratory strains or those studied under clinical settings at temperatures where luminescence may not be produced. With environmental isolates and previously characterized species, the use of cooler temperatures (10–20∞C) for growth and examination, utilization of conditioned media, inducers and luciferase substrates (Fidopiastis et al., 1999), and the application of probes for luxA and other lux genes (Wimpee et al., 1991) will undoubtedly reveal many more types of bacteria with the ability to produce light. From the perspective of 16S rRNA sequencebased phylogeny, the luminous bactera are representative members of the g-Proteobacteria, with luminous species in four genera (Vibrio, Photobacterium, Photorhabdus and Shewanella) within three families (Vibrionaceae, Enterobacteriaceae and Alteromonadaceae; Fig. 2). Genera containing species or strains of luminous bacteria are a small fraction of the162 genera in 20 families of the g-Proteobacteria (see Bergey’s Manual of Systematic Bacteriology, May 2001). Most of the luminous species are members of the Vibrionaceae (genera Vibrio and Photobacterium), which also contain many nonluminous species. Diverging from lineages within the Vibrionaceae are several uncultured luminous symbionts of anomalopid (flashlight) and ceratioid (deep-sea) anglerfish (Fig. 2). The anomalopid and ceratioid symbionts form separate monophyletic groups, and these symbionts are sufficiently divergent from known luminous bacteria to suggest they represent new species or genera within the Vibrionaceae (Haygood, 1990; Haygood, 1993a; Haygood and Distel, 1993b). The intermingling of luminous and nonluminous species in the Vibrionaceae contrasts with the phylogenetic separateness of the luminous species within the Enterobacteriaceae and Alteromonadaceae (Fig. 2). Within these latter two families, the luminous species occur on branches that appear distal to other species. Placement of Photorhabdus in the Enterobacteriaceae, though generally accepted, is controversial (Janse and Smits, 1990; Rainey et al., 1995), however. Characteristics of Photorhabdus species not typical of members of the Enterobacteriaceae include luminescence, synthesis of yellow and red pigments and the inability to reduce nitrate (Farmer et al., 1989; Forst and Nealson, 1996).

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CHAPTER 1.27

Table 1. Luminous bacteria. Speciesa Marine Vibrio fischeri

Temperate coastel seawater

Bioluminescent symbiosis

Monocentrid fish, certain sepiolid squids

— — — — —

Nealson et al., 1993 Oliver et al., 1986

Temperate to tropical coastal seawater, sediment

logei

Unnamed

Coastal cold seawater, sediment, Arctic and Mediterranean Coastal seawater Seawater, surfaces of shrimp Tissue lesions of Atlantic salmon Coastal seawater, Persian Gulf Human blood and tissue, United States Not yet cultured

Anomalopid fish

Unnamed

Not yet cultured

Ceratioid fish

splendidus vulnificus

Photobacterium angustum leiognathi

phosphoreum

Shewanella hanedai woodyi Brackish/Estuarine Vibrio cholerae Terrestrial Photorhabdus luminescens temperata asymbiotica

Seawater and fish intestines, Sea of Cortez Coastal temperate to tropical seawater leiognathid fish Coastal and pelagic cold to temperate seawater

Selected references

Boettcher and Ruby, 1990 Fitzgerald, 1977 Lee and Ruby, 1992 Reichelt and Baumann, 1973 Ruby and Nealson, 1976 Ruby and Nealson, 1978 O’Brien and Sixemore, 1979 Reichelt and Baumann, 1973 Ruby and Nealson, 1978 Yetinson and Shilo, 1979 Bang et al., 1978 Baross et al., 1978 Fidiopastis et al., 1998 Ortiz-Conde et al., 1989 Yang et al., 1983 Fidiopastis et al., 1999

harveyi

mediterraneab orientalis salmonicida

a

Representative habitats

—

Certain sepiolid squids

— Acropomatid, apogonid, Certain loligiroid squids Opisthoproctid, chlorophthalmid, trachichthyid, morid, macrourid, steindachnerid fish

Hygood, 1990 Wolfe and Haygood, 1991 Haygood and Distel, 1993 Haygood et al., 1992 Kita-Tsukamoto et al. (in prep.) Fukasawa and Dunlap, 1986 Fukasawa et al., 1998 Herring and Morin, 1978 Reichelt et al., 1977 Herring and Morin, 1978 Haygood, 1993 Ruby and Morin, 1978 Wimpee et al., 1991

Cold seawater and sediment Seawater and squid ink, Alboran Sea

— —

Jensen et al., 1980 Makemson et al., 1997

Temperate to tropical estuaries, bays coastal seawater

—

Palmer and Colwell, 1989 Ramaiah et al., 2000

Insect larvae infected with heterorhabditid nematodes Insect larvae infected with heterorhabditid nematodes Human skin lesions United States and Australia

—c

Boemare et al., 1993 Fischer-Le Saux et al., 1999 Fischer-Le Saux et al., 1999

—c —

Farmer et al., 1989 Fischer-Le Saux et al., 1999 Peel et al., 1999

Luminous strains. For additional information, see Baumann and Baumann (1981); Farmer and Hickman-Brenner (1992); Hastings and Nealson (1981); and Nealson and Hastings (1992). b Ability of this species to luminesce is not well established. c Symbiotic with entomopathogenic nematodes; on anatomical and behavioral grounds not considered here to be equivalent to bioluminescent symbiosis in fishes and squids.

CHAPTER 1.27

Luminous Bacteria

867

Akat/symb Kryp/symb Pste/symb Ppal/symb

Enterobacteriaceae Vcho

Plumlau Plumakh Plumlum Pasy Ptern Xnem 992 Xjap Xbed Xbov Smar

Vang Vvul Vhar Vibrionaceae VfisVsal Vflu Vpel MJ1/Vfis Vlog Vpar Vmed Vori Vgaz 991 Phis 810 Ppro Plei 999 810 638 PL721/Plei GB1/Pang 996 Pang Ppho 744 Pili Og61/Ppho 1000 Vspl

Ecar Vhol Ecol

Scos

Yent

Mjoh/symb

Shan Swoo

Ccou/symb

Sben Scol Alteromonadaceae

Salg Sput

0.01 Mmar

Fig. 2. 16S rDNA-based phylogenetic tree of luminous bacteria. Luminous species and strains are in boldface. Names are abbreviated as the first letter of the genus and the first three letters of species of the bacterium or its symbiotic host, along with the first three letters of the subspecies, where appropriate. See accession numbers as indicated for references. Included in the tree for comparison are sequences of species and strains related to luminous bacteria and sequences of uncultured symbiotic luminous bacteria. The neighbor-joining (NJ; Saitou and Nei, 1987) tree was constructed based on 1,163 unambiguously aligned positions. CLUSTAL W program (ver. 1.60; Thompson et al., 1994) was used for alignment of sequences and realigned manually using MacClade 3.05 (Maddison and Maddison, 1992). The NJ tree was developed from the distance matrix calculated by the algorithm of the Kimura two-parameter model (Kimura, 1980). Bootstrap analysis was done with 1,000 replicates. The scale bar represents 0.01 nucleotide substitutions per position. Alteromonadaceae: Moritella marina (T [type strain]), X82142; Shewanella algae (T), AF005249; S. benthica (T), X82131; S. colwelliana (T), AF170794; S. hanedai (T), X82132; S. woodyi (T), AF003549; and S. putrefaciens (T), X81623. Enterobacteriaceae: Erwinia carotovora subsp. carotovora (T), M59149; Escherichia coli, J01859; Photorhabdus asymbiotica (T), Z76755; P. luminescens subsp. akhurstii (T), AJ007359; P. luminescens subsp. laumondii (T), AJ007404; P. luminescens subsp. luminescens (T), X82248; P. temperata (T), AJ007405; Serratia marcescens (T), M59160; Xenorhabdus beddingii (T), X82254; X. bovienii (T), X82252; X. japonicus (T), Z76739; X. nematophilus (T), X82251; and Yersinia enterocolitica (T), M59292. Vibrionaceae: Photobacterium angustum (T), X74685; P. angustum GB-1 (K. Kita-Tsukamoto et al., manuscript in preparation); P. histaminum, D25308; P. iliopiscarium (T), AB000278; P. leiognathi (T), X74686; P. leiognathi PL-721, Z21730; P. phosphoreum (T), D25310; P. phosphoreum Og61, Z19107; P. profundum, AB003191; Salinivibrio costicola (T), X74699; Vibrio anguillarum (T), X16895; V. cholerae, X74694; V. fischeri (T), X74702; V. fischeri MJ-1, Z21729; V. fluvialis (T), X74703; V. gazogenes (T), X74705; V. harveyi (T), X74706; V. hollisae (T), X74707; V. logei, X74708; V. mediterranei (T), X74710; V. orientalis (T), X74719; V. parahaemolyticus (T), X74720; V. pelagius (T), X74722; V. salmonicida (T), X70643; V. splendidus (T), X74724; and V. vulnificus (T), X74726. Uncultured symbiotic luminous bacteria: Anomalops katoptron symbiont, Z19081; Cryptosaras couesi symbiont, Z19106; Kryptophanaron alfredi symbiont, Z19003; Melanocetus johnsoni symbiont, Z19105; Photoblepharon palpebratus symbiont, Z19085; and Photoblepharon steinetzi symbiont, Z19080.

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Nonetheless, the separateness of the luminous species in these two families suggests a separate and relatively recent acquisition of lux genes by Photorhabdus (Forst et al., 1997) and luminous Shewanella (discussed below).

Habitats and Ecology of Luminous Bacteria Marine Luminous bacteria are globally distributed in the marine environment (Table 1) and can be isolated from seawater, sediment and suspended particulates. They also colonize marine animals as saprophytes, commensal enteric symbionts and parasites (Hastings and Nealson, 1981; Meighen and Dunlap, 1993; Makemson et al., 1997), and certain of them establish bioluminescent symbiosis with marine fish and squids (Dunlap and Greenberg, 1991b; Nealson and Hastings, 1992; Haygood, 1993a; Ruby, 1996). In seawater, numbers of luminous bacteria generally are low (from 0.01 to up to 40 cells per ml of seawater; Nealson and Hastings, 1992). In association with animals, however, luminous bacteria can attain very high numbers (up to 1011 cells per ml in symbiotic habitats; Ruby and Nealson, 1976; Dunlap, 1984; Nealson and Hastings, 1992). The very high numbers of luminous bacteria in saprophytic, commensal, parasitic and symbiotic habitats indicates the potential of these habitats to make substantial contributions to the density and distribution of luminous bacteria in seawater, sediments and marine snow (Reichelt et al., 1977; O’Brien and Sizemore, 1979; Ruby and Morin, 1979; Haygood et al., 1984; Nealson et al., 1984; Ramesh et al., 1987; Ruby and Lee, 1998), which in turn serve as environmental sources of these bacteria for re-colonization of animals (Nealson and Hastings, 1992). Except in bioluminescent symbiosis, which is specific to certain luminous bacteria, the luminous species coexist in these habitats with nonluminous bacteria. Unlike marine animals, marine algae apparently are not commonly colonized by luminous bacteria. Only one luminous bacterium with the ability to digest agar has been reported, a strain of V. harveyi (Fukasawa et al., 1987). Although agar digestion is often observed among Vibrio spp. and other marine bacteria, searches for other luminous bacteria with this trait, including extensive examination of surfaces of marine algae, have not yet revealed other luminous strains. One can speculate that a rare bacteriophage-mediated transduction between Vibrio spp. (Baross et al., 1978a) might have been the mechanism by which the strain of V. harveyi acquired

CHAPTER 1.27

genes for agar hydrolysis. Regardless, the surfaces of marine algae are an additional habitat exploited, though apparently rarely, by luminous bacteria and shared with nonluminous forms. The distributions and numbers of individual species of luminous bacteria correlate with certain environmental factors (Baumann and Baumann, 1981; Hastings and Nealson, 1981). Primary among these factors are temperature and depth (Ruby and Nealson, 1978b; Yetinson and Shilo, 1979; Ruby et al., 1980), salinity (Yetinson and Shilo, 1979; Feldman and Buck, 1984), nutrient limitation and sensitivity to photooxidation (Shilo and Yetinson, 1980). Temperature, along with being an important environmental factor, can influence whether luminous bacteria from environmental samples are detected. For example, Shewanella hanedai and Vibrio logei, which are psychrotrophic, grow and produce light at low temperature (e.g., 4∞C to 15∞C, and grow but do not produce light at room temperature (24∞C). Therefore, incubation of platings of environmental samples at the lower temperatures may reveal the presence of other psychrotropic luminous species. Temperature relationships would appear to be species-specific, however. For example, S. woodyi (found in squid ink and seawater in the Alboran Sea near Gibraltar; Makemson et al., 1997) and V. fischeri, species closely related to S. hanedai and V. logei, respectively, grow and produce light at room temperature. Studies of the distibution and density of luminous bacteria in the marine environment traditionally have used visual observation of luminescence to identify these bacteria. However, the presence of lux genes in bacteria that do not produce light in culture and the physiological crypticity of luminescence in some species (Boettcher and Ruby, 1990; Fidopiastis et al., 1999) reveal that luminous bacteria are more numerous and diverse than identified by the luminescence phenotype. Enzyme assay and antibody methods previously have been used to detect luciferase in several nonluminous Vibrio spp. (Nealson and Walton, 1978b; Makemson and Hastings, 1986b; Kou and Makemson, 1988), and luxA-based DNA probes from various seawater samples have been used to identify lux gene-containing bacteria not producing light in culture (Potrikus et al., 1984; Palmer and Colwell, 1991; Lee and Ruby, 1992; Wimpee et al., 1991; Ramaiah et al., 2000). The efficacy of species- and group-specific luxA-based probes for the identification of environmental isolates of luminous bacteria has been demonstrated for two species, Photobacterium phosphoreum from the Black Sea and Vibrio splendidus from coastal waters of Kuwait (Wimpee et al., 1991; Nealson et al., 1993).

CHAPTER 1.27

Freshwater Knowledge of luminous bacteria in freshwater environments is limited to reports that luminous V. cholerae exist in freshwater and infect freshwater crustaceans. Luminous strains of V. cholerae have been isolated from freshwater and brackish estuarine waters in various locations (Desmarchelier and Reichelt, 1981; West and Lee, 1982; West et al., 1983; Palmer and Colwell, 1991; Ramaiah et al., 2000; Table 1). The first such isolation, in 1893, apparently was by F. Kutscher from the Elbe River in Germany (Harvey, 1952). Then called “Vibrio albensis,” that strain later was synonymized with V. cholerae (Reichelt et al., 1976). With respect to infecting freshwater animals, Thulis and Bernard in 1786 described the luminescence of a freshwater crustacean (possibly the common amphipod Gammarus pulex, which apparently was infected with luminous bacteria) from a river in southern France (Harvey, 1957). Yasaki (1927) reported the isolation of luminous bacteria from intensely luminous specimens of the freshwater shrimp, Xiphocaridina compressa, in Lake Suwa, Japan. Initially characterized as Microspira phosphoreum, the bacterium was later redescribed as Vibrio yasakii (Majima, 1931). More recently, a bacterium responsible for this “light disease of shrimp” was isolated from freshwater shrimp in Lake Biwa, Japan, and identified as non-O1 V. cholerae (Shimada et al., 1995). Nonluminous V. cholerae also are associated with disease in freshwater crustaceans (Thune et al., 1991).

Terrestrial Luminous bacteria in the terrestrial environment have been noticed mostly as parasites of insects that cause the infected animal to luminesce. Observations of luminous midges, caterpillars, mole-crickets, mayflies and ants, among other infected insects, have been reported from the 1700s into modern times (Harvey, 1952; Haneda, 1950). As described and summarized by Harvey (Harvey, 1952; Harvey, 1957), other early reports of terrestrial luminescence attributable to luminous bacteria include luminous mutton, veal, eggs of chickens and lizards, human corpses and battlefield wounds. Many, and perhaps all, of the observations of luminous insects result from colonization by members of the genus Photorhabdus, of which three species are currently described, P. luminescens, P. temperata and P. asymbiotica (Fischer-Le Saux et al., 1999; Table 1). Photorhabdus luminescens and P. temperata occur as the mutualistic symbionts of entomopathogenic nematodes (commonly found in

Luminous Bacteria

869

soil) of the family Heterorhaditidae (Akhurst and Dunphy, 1993; Forst and Nealson, 1996; Forst et al., 1997). They are carried in the intestine of the infective juvenile stage of the nematode and participate in a lethal infection of insect larvae. When the nematode enters the insect, via the digestive tract or other openings, and penetrates the insect’s hemocele, the bacteria are released into the hemolymph, where they use its constituents for growth. The bacteria elaborate a variety of extracellular enzymes that presumably break down macromolecules of the hemolymph. Proliferation of the bacteria leads to death of the insect, and its carcass becomes luminous. The bacteria also produce various extracellular and cell surface-associated factors pathogenic for the insect, as well as bacteriocins and hydroxystilbene and anthraquinone antibiotics, which apparently inhibit the growth of other microorganisms in the insect cadaver (Akurst, 1982). Crystalline protein inclusion bodies of unknown function are also produced (Bintrim and Ensign, 1998). The nematodes feed on the bacteria or products of bacterial degradation of the hemolymph enabling them to develop and sexually reproduce (Boemare et al., 1997; Forst et al., 1997). Completion of the nematode life cycle involves reassociation with the bacteria and the emergence from the insect cadaver of the nonfeeding infective juveniles, carrying the bacteria in their intestines. Cells of P. luminescens presumably are present in soil, but association with the nematode apparently is important for their survival and dissemination. Luminescence of the infected insect larva might function to attract nocturnally active animals to feed on the glowing carcass, thereby increasing the opportunities for the bacterium and the nematode to be disseminated. However, luminescence is not required for successful symbiosis with the nematode; not all strains of P. luminescens produce luminescence (Akhurst and Boemare, 1986; Forst and Nealson, 1996). Furthermore, bacteria in the genus Xenorhabdus, which are symbiotic with entomopathogenic nematodes in the family Steinernematidae, are ecologically very similar to Photorhabdus, except that they do not produce light (Akhurst and Dunphy, 1993). The similarities between the lifestyles and activities of Photorhabdus and Xenorhabdus are postulated to be a case of ecological convergence (Forst and Nealson, 1996). Human clinical infections have yielded P. asymbiotica, introduced apparently by spider and insect bites (Farmer et al., 1989; Peel et al., 1999). Luminous battlefield wounds are intriguing because luminescence apparently is a sign that the wound will heal well (Harvey, 1957). Indeed, luminous bacteria will grow and produce light on living mammalian tissue (Johnson, 1988). Perhaps antibiotic-producing P. lumine-

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scens or P. temperata promoted wound healing by preventing the growth of putrefying, pathogenic bacteria. On the other hand, the human pathogenicity of P. asymbiotica suggests that this species might have killed rather than healed if introduced into wounds. The recent description of P. asymbiotica and P. temperata, and the presence of genetically distinct subspecies within P. luminescens and P. temperata (Fischer-Le Saux et al., 1999; Fig. 2) indicate that additional diversity, possibly at the species level, may exist in this genus. Along with terrestrial Photorhabdus species, marine luminous bacteria might have been responsible for some of the early reports of luminous meats and eggs, especially if brine was used in their preparation or they otherwise were exposed to seawater. Haneda (1950), following the observation by Molisch (1925) of luminous bacteria growing on beef, demonstrated that luminous bacteria could be isolated from certain samples of beef, pork and chicken meat. These meats might have contained enough salt to support the growth of marine forms, and Haneda cultured the bacteria in media containing 0.5% salt. However, whether these bacteria were terrestrial (i. e., Photorhabdus), freshwater (i. e., V. cholerae), or marine in origin apparently is not known.

Parasitism of Marine Invertebrates Most of the commonly encountered marine luminous bacteria are not known to be highly invasive or virulent in animals. Many or perhaps all luminous species, however, can act as opportunistic pathogens upon entering an animal’s body through lesions resulting from injury or stress. First noted in marine animals apparently by Viviani in 1805 (Harvey, 1957), infections of marine crustaceans by luminous bacteria are common, causing the infected animal to luminesce (Giard, 1889a; Giard and Billet, 1889b; Inman, 1926; Hastings and Nealson, 1981). Luminous bacteria inhabit the gut tract and colonize external surfaces of marine crustaceans (Inman, 1926; Baross et al., 1978b; O’Brien and Sizemore, 1979; Lavilla-Pitogo et al., 1992); many are chitinolytic (Spencer, 1961; Baumann and Schubert, 1984a). The bacteria enter the hemocele of the animal through lesions in the gut or carapace, developing luminescence and killing the animal within a few days. The species of luminous bacteria infecting isopods and amphipods commonly encountered in coastal environments have not been identified in recent times, but they exhibit characters consistent with members of the genera Vibrio and Photobacterium (Hastings and Nealson, 1981; P. Dunlap, unpublished observation). Nonluminous bacteria undoubt-

CHAPTER 1.27

edly cause similar infections that go unnoticed due to the lack of light production. As opportunistic pathogens of marine crustaceans, luminous bacteria have had a profoundly deleterious effect on commercial prawn mariculture. The development of intensive monoculture of Penaeus monodon, the giant tiger prawn, and other penaeids during the 1980s led to a dramatic increase in disease and death of the animals due to luminous bacteria. Shrimp hatchery rearing ponds can become heavily infested with luminous bacteria, with shrimp larvae developing “luminescent vibriosis,” a pathogenic state responsible for massive mortalities. The problem continues in grow-out ponds, where the infection localizes to the hepatopancreas in juveniles, limiting the growth of the animals and further increasing losses to mortality (Lavilla-Pitogo and de la Peñ a, 1998). Primarily responsible are strains of V. harveyi, though other luminous and nonluminous vibrios have been identified (Lavilla-Pitogo et al., 1990; Karunasagar et al., 1994; Lavilla-Pitogo and de la Peñ a, 1998; Leano et al., 1998).

Parasitism of Vertebrates In contrast to the situation with marine invertebrates, luminous bacteria apparently only rarely infect vertebrate animals. The ability of P. asymbiotica to infect humans has been mentioned above. Vibrio harveyi has been identifed in fish disease, and recently, V. salmonicida (a pathogen of salmonids and cod) has been shown to produce light under certain conditions (Fidopiastis et al., 1999). Clinical strains of Vibrio vulnificus and V. cholerae typically are nonluminous, but luminous strains of V. vulnificus have been isolated from dead humans (Oliver et al., 1986), and luminous strains of V. cholerae have been isolated from humans suffering from cholera (Jermoljewa, 1926). Furthermore, Weleminsky (1895) demonstrated that a nonluminous clinical isolate of V. cholerae developed luminescence apparently by passage through another animal. Vibrio cholerae strains that are luminous or that contain the luxA gene are present in relatively high percentages in freshwater and estuarine environments (West and Lee, 1982; West et al., 1983; Palmer and Colwell, 1991; Ramaiah et al., 2000). The lightproducing and luxA gene-containing strains are the non-O1 type of V. cholerae (Palmer and Colwell, 1991; Ramaiah et al., 2000).

Bioluminescent Symbiosis One of the most remarkable attributes of luminous bacteria is the ability of certain species to

CHAPTER 1.27

establish luminescence-based symbiotic associations called “bioluminescent symbiosis” with marine animals. These associations have been found in certain teleost fish, some loliginid and sepiolid squids, and possibly in pyrosomes and salps. The treatise by Buchner (1965) and the review by Herring and Morin (1978b) provide comprehensive access to early literature. For pyrosomes and salps, bioluminescent symbiosis with luminous bacteria is controversial (Harvey, 1952; Buchner, 1965). Pyrosome zooids bear a pair of simple photophores containing intracellular bacteroids, but the involvement of bacteria in pyrosome luminescence has been both discounted and supported (Galt, 1978; Herring, 1978a; Mackie and Bone, 1978; Haygood, 1993a). Although the bacteroids have not been cultured, the presence of bacterial luciferase in photophores is consistent with a bacterial origin for pyrosome luminescence (Leisman et al., 1980). In myctophid and stomiiform fishes, a similar proposal that the luminescence of photophores is due to the presence of symbiotic luminous bacteria (Foran, 1991) was shown conclusively to be invalid (Haygood et al., 1994). In bioluminescent symbiosis of fishes and squids with luminous bacteria (Table 1), the host animal bears one or a pair of specialized glandlike tissues, called “light organs,” which house a pure culture of the species-specific symbiotic bacterium. Accessory structures associated with the light organ, i.e. lens, reflector, and lightabsorbing shutters and barriers, control, direct and focus the light the bacteria produce. The host animal uses the bacterial light in luminescence displays associated with various behaviors, including predator avoidance by counterillumination and flashing, sex-specific signaling, attracting or locating prey, and orienting in dark and dimly lighted environments (Hastings, 1971; Morin et al., 1975; Nealson and Hastings, 1979b; McFall-Ngai and Dunlap, 1983; McFall-Ngai and Montgomery, 1990; McFall-Ngai and Morin, 1991a). In fishes, the light organs are internal, associated with the gut tract, or external, located below the eye (subocular light organ), in the lower jaw (mandibular light organ) or at the terminus of an elongated dorsal fin ray (escal light organ), whereas in squids, they are found as bilobed organs ventrally within the mantle cavity, associated with the ink sac (Herring, 1977; Hastings and Nealson, 1981; Haygood, 1993a; McFall-Ngai and Ruby, 1991b). Bioluminescent symbiosis, owing to the specificity between host and symbiont, contrasts with other associations of luminous bacteria with animals; the saprophytic, commensal and parasitic associations are nonspecific and often involve assemblages of luminous and nonluminous bacteria (Harvey, 1940; Nealson and Hastings, 1992). Fur-

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thermore, a trend toward greater integration between symbiont and host can be envisioned in bioluminescent symbiosis, with certain animals colonized by facultatively symbiotic (i.e., culturable) bacteria and others harboring obligately symbiotic (i.e., not yet cultured) bacteria (Haygood, 1993a). Bioluminescent symbiosis appears to be a unique kind of symbiosis; the bacterial metabolic product needed by the host animal is light, used in bioluminescence displays, rather than a nutrient needed for host development or growth (Claes and Dunlap, 2000). Four species of luminous bacteria, V. fischeri, V. logei, P. leiognathi and P. phosphoreum, have been identified in bioluminescent symbiosis (Table 1). Vibrio fischeri and P. leiognathi colonize light organs of both fish and squids (Boettcher and Ruby, 1990; Fukasawa et al., 1986; Fukasawa et al., 1988; Reichelt et al., 1977), whereas P. phosphoreum so far has been found in association only with fish (Herring and Morin, 1978b; Hastings and Nealson, 1981). Vibrio logei was identified recently as the predominant symbiont of the sepiolid squids Sepiola affinis and Sepiola robusta (Fidopiastis et al., 1998). Two other groups of fishes, the flashlight fish (family Anomalopidae) and deep-sea anglerfish (suborder Ceratioidei) bear light organs with symbiotic luminous bacteria that so far have not been cultured. The anomalopid symbionts, based on analysis of the luxA and 16S rRNA genes, are likely to be members of the genus Vibrio, and different genera of the fish harbor bacteria that differ at greater than the strain level (Haygood, 1990; Wolfe and Haygood, 1991). The results of 16S rRNA gene sequence analysis of the bacterial symbionts of two ceratioids (representing different families of anglerfish) group these bacteria with other marine enterics phylogenetically similar to Photobacterium and Vibrio and suggest that these may be new bacterial species in each fish (Haygood and Distel, 1993b).

Symbiont-Host Specificity Despite the presence of various different species of luminous bacteria in the habitats of animals that form bioluminescent symbiosis, these associations are highly specific. Members of a given family of fishes and of squids consistently harbor the same species of symbiont (Fitzgerald, 1977; Reichelt et al., 1977; Ruby and Morin, 1978a; Ruby and Nealson, 1976; Fukasawa et al., 1986; Hastings and Nealson, 1981; Ruby, 1996). Various selective pressures, alone or in combination might account for this specificity. Physiological conditions of the light organ, which derives from host biology and light organ anatomy, may be important. Light-organ osmolarity, oxygen and iron levels, and types of nutrients presum-

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ably interface with the individual physiological capabilities of different symbiotic luminous species and strains, promoting adaptively high levels of luminescence and competitive growth dominance of one type over another (Ruby and Nealson, 1976; Ruby and Nealson, 1977; Nealson, 1979a; Dunlap, 1985a; Haygood and Nealson, 1985a; Haygood, 1993a; Hastings et al., 1987; Graf and Ruby, 1998; Lee and Ruby, 1994a; Visick et al., 2000). Specific recognition and other exclusion mechanisms (Ruby, 1996) also may play a major role. Local abundance of the host also may contribute to specificity (Hastings and Nealson, 1981; Ruby and Lee, 1998). Along with host-related factors, temperature, as an environmental factor, might play a significant role in host-symbiont specificity. A loose concordance is seen between the temperature of the host’s habitat and the temperature sensitivities of the symbiotic bacteria. Fishes dwelling in temperate and tropical shallow waters harbor the more mesophilic species V. fischeri or P. leiognathi, whereas fishes dwelling in cold, deeper waters tend to harbor the more psychrotrophic species P. phosphoreum (Hastings and Nealson, 1981). Indeed, temperature reveals an exception to the pattern of host-symbiont specificity. The closely related species V. logei and V. fischeri can colonize the same species of sepiolid squid, Sepiola affinis and Sepiola robusta, forming mixed symbiotic cultures. Whereas lower temperatures favor the more psychrotrophic V. logei, warmer temperatures favor the more mesophilic V. fischeri (Fidopiastis et al., 1998; Nishiguchi, 2000).

Symbiont Transmission In the few cases studied, squids and fishes have been found to acquire their symbiotic luminous bacteria by horizontal transfer. Best documented is the sepiolid squid Euprymna scolopes, hatchlings of which carry no V. fischeri cells or other bacteria in their nascent light organs. Soon after hatching, the animal picks up its symbiotic bacterium from seawater, establishing bioluminescent symbiosis (Wei and Young, 1989; McFallNgai and Ruby, 1991b). Symbiont motility is required for this process (Graf et al., 1994). Nascent light organs of juvenile Siphamia versicolor (family Apogonidae) at the early larval stage lack bacteria but contain them later in development, consistent with acquisition of the bacteria from the seawater (Haneda, 1965; Leis and Bullock, 1986). Recently, Wada et al. (1999) provided evidence for horizontal transfer of P. leiognathi to juvenile Leiognathus nuchalis (family Leiognathidae). For anomalopid (flashlight) fish, no evidence was found that the symbiotic bacteria are associated with gonads or eggs, con-

CHAPTER 1.27

sistent with horizontal transfer in this group as well (Haygood, 1993a).

Symbiont Contributions to Host Survival, Growth, and Development Bioluminescent symbiosis appears to be a special class of symbiosis, one in which the primary metabolic contribution the symbiotic bacteria make to the host is light. In most bacterial associations with animals and plants, the host is dependent nutritionally on its symbiotic bacteria, via bacterial fixation of carbon or nitrogen, the activity of bacterial extracellular degradative enzymes, such as cellulases, or bacterial provision of vitamins or other essential nutrients (Douglas, 1995). As a consequence, absence of the symbiotic bacteria can have a profound influence on the survival, growth and development of the host. In contrast, the sepiolid squid E. scolopes cultured aposymbiotically from hatching to reproductive adulthood survived, grew and developed equally as well as animals colonized by V. fischeri (Claes and Dunlap, 2000). These observations indicate that V. fischeri apparently makes no major nutritional contribution to the animal. The metabolic dependency of E. scolopes on V. fischeri therefore seems limited to light production. Selection for the association presumably is ecological, with the squid’s ability to counterilluminate using light produced by V. fischeri (Singley, 1983; McFall-Ngai and Montgomery, 1990) playing an important role in survival. Whether a similar lack of nutritional dependency of the host on its symbiotic bacteria characterizes other bioluminescent symbioses remains to be determined.

Influence of Symbionts on Light-Organ Morphogenesis Much progress in understanding host-symbiont relationships in bioluminescent symbiosis has developed from studies of the sepiolid squid E. scolopes. The animal maintains a speciesspecific (and strain-specific) association with V. fischeri, harboring the bacteria extracellularly in diverticulated epithelial tubules comprising the core of the bilobed ventral light organ. Associated with the light organ are accessory tissues, specifically the ink sac, and the reflectors and lens, which control and direct the light produced by V. fischeri. The tubules lead into a ciliated duct that connects each lobe of the light organ to the mantle cavity (McFall-Ngai and Ruby, 1991b; Ruby, 1996; McFall-Ngai, 1999). Analysis of the colonization process in hatchling juvenile E. scolopes demonstrates a role for the symbiotic bacterium in morphological changes in the light organ. The nascent, rudimentary light organs in hatchlings bear a pair of

CHAPTER 1.27

lateral ciliated epithelial appendages (CEAs) and contain a pair of three simple sac-like epithelial tubules embedded in the undifferentiated accessory tissues. The proximal portions of these tubules are ciliated and directly connect to the mantle cavity via a lateral pore. Colonization of the epithelial tubules, which is facilitated by ciliary beating of the CEAs, occurs through these lateral pores, which later coalesce, with the formation of a ciliated duct for each light organ lobe. Colonization triggers regression of the CEAs within approximately 4 days. Other morphological changes include alterations in the epithelial cells of the distal portions of the light organ tubules, which develop a dense microvillous brush border (McFall-Ngai and Ruby, 1991b; Montgomery and McFall-Ngai, 1993; Montgomery and McFall-Ngai, 1994; Doino and McFall-Ngai, 1995; Ruby, 1996; Lamarcq and McFall-Ngai, 1998; McFall-Ngai, 1999). Presence of the bacteria, however, is not necessary for overall development of the light organ and its accessory tissues, which proceed normally in aposymbiotic animals (Claes and Dunlap, 2000). In possible contrast to light organ development in E. scolopes, light organs of the monocentrid fish, M. japonicus, had not developed by day 21 in larvae from artificially fertilized eggs (Yamada et al., 1979), suggesting that acquisition of V. fischeri may be necessary to initiate the light organ developmental program. Whether development of the light organ in aposymbiotic juvenile leiognathid fish requires colonization by P. leiognathi is not yet known (Wada et al., 1999).

Host Contribution to Symbiont Dissemination Bioluminescent symbiosis is likely to have a significant impact on the density and distribution of the symbiotic bacteria in seawater. Growth of the bacterial population in the light organ leads to the continual or diurnal release of bacterial cells into the environment (Dunlap, 1984; Haygood et al., 1984; Nealson et al., 1984; Lee and Ruby, 1994b; Boettcher et al., 1995). The cells are released either directly into seawater, as in monocentrid fish and sepiolid squids, or indirectly via the gut tract, as in leiognathid fish. Estimates of growth rates for the bacteria indicate the population doubles once to a few times per day (Dunlap, 1984; Haygood et al., 1984; Lee and Ruby, 1994b), so each adult host may release as many as 107 to 108 symbiont cells per day. This release, which has the potential of dispersing the bacteria into other habitats they colonize (Nealson et al., 1984), may be essential for re-initiation of the association with the next generation of the host animal (Ruby and Lee, 1998).

Luminous Bacteria

873

Physiological Control of Luminescence Growth conditions can strongly influence the amount of light produced by luminous bacteria in laboratory culture. Oxygen, amino acids, glucose, iron and osmolarity have distinct effects, depending on the species studied (Harvey, 1952; Nealson and Hastings, 1977b; Makemson and Hastings, 1982; Haygood and Nealson, 1985a; Hastings et al., 1987; Dunlap, 1991a). Those factors that stimulate growth rate, such as readily metabolized carbohydrates, tend to decrease light production and luciferase synthesis. They do so presumably by causing oxygen and reducing power (FMNH2) to be directed away from luciferase (McElroy and Seliger, 1962; Coffey, 1967) and by indirectly or directly influencing lux gene expression (Dunlap and Greenberg, 1985b; Dunlap, 2000). Conversely, factors that restrict growth rate, such as limitation for iron, tend to stimulate the synthesis and activity of luciferase (Hastings and Nealson, 1977; Haygood and Nealson, 1985a; Hastings et al., 1987; Dunlap, 1991a). The mechanisms by which these factors operate, however, are not well understood (Haygood and Nealson, 1985b; Dunlap, 1992a; 1992b), indicating that much remains to be learned about the interplay between growth physiology of the cell and regulatory elements controlling lux gene expression.

Amino Acids, Catabolite Repression, and Control by cAMP The amino acid arginine and certain structurally and metabolically related compounds can stimulate luminescence in V. harveyi growing in minimal medium (Coffey, 1967; Nealson et al., 1970; Hastings and Nealson, 1977). The mechanism for this activity remains unknown. Conversely, yet equally intriguing, mixtures of amino acids can transiently and in a dose-dependent manner block the increase in light production of inducing cultures of P. leiognathi (P. Dunlap, unpublished observation). Catabolism of the amino acids might account for this temporary repression of luminescence induction. Catabolite repression of luminescence generally is attributed to effects on levels of 3¢, 5¢-cyclic AMP (cAMP) and cAMP receptor protein (CRP; Nealson et al., 1972; Meighen and Dunlap, 1993; Dunlap, 1997). Different species of luminous bacteria, however, respond in different ways. In V. harveyi, catabolite repression by glucose in batch culture is permanent and is reversed by addition of cAMP (Nealson et al., 1972), whereas glucose repression of luminescence in V. fischeri is temporary, is not reversed

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by addition of cAMP, and is eliminated by prior growth in the presence of glucose (Ruby and Nealson, 1976). Complicating these differences from studies in batch culture are studies of V. fischeri grown in phosphate-limited chemostat culture; glucose repression of luminescence then is permanent and reversible by cAMP (Friedrich and Greenberg, 1983). A further complication for studies of cAMP-control of luminescence in V. fischeri is the presence in this species of a novel, exceptionally potent periplasmic cyclic nucleotide phosphodiesterase specific for extracellular 3¢, 5¢-cyclic nucleotides; activity of the enzyme enables cells to grow on exogenously supplied cAMP as a sole source of carbon and energy, nitrogen and phosphorus (Dunlap et al., 1992d; Dunlap and Callahan, 1993; Callahan et al., 1995). Regardless of the differences in catabolite repression, mutants of V. harveyi and V. fischeri apparently defective in adenylate cyclase and unable to produce light in the absence of added cAMP have been isolated and characterized (Ulitzur and Yashphe, 1975; Dunlap, 1989a). Furthermore, CRP from V. harveyi has been purified and shown to be immunologically and functionally homologous to CRP of Escherichia coli (Chen et al., 1985), and the cya and crp genes of V. fischeri have been cloned and found to be highly similar in deduced amino acid residue sequence to E. coli cya and crp genes (P. Dunlap et al., unpublished observation). Consistent with these observations, the regions upstream of the luminescence operons of V. harveyi and V. fischeri contain a CRP binding site (Engebrecht and Silverman, 1987; Devine et al., 1988a; Miyamoto et al., 1988b). Studies with V. fischeri and with E. coli carrying the V. fischeri luminescence system indicate that a major effect of cAMP-CRP is to activate the production of LuxR, the luminescence operon transcriptional activator (see below; Dunlap and Greenberg, 1985b; Dunlap and Greenberg, 1988; Dunlap and Kuo, 1992c; Shadel et al., 1990a), although other important lux regulatory effects have also been described (Shadel and Baldwin, 1991; Shadel and Baldwin, 1992a; Shadel and Baldwin, 1992b). Regardless, control of the luminescence system by cAMP-CRP in V. fischeri demonstrates the integration of luminescence with cellular metabolism and suggests that activity of the luminescence system is part of the cellular response to stresses associated with nutrient limitation and decreasing growth rate. Consistent with that view, a heat-shock protein (GroESL) and a repressor of DNA repair (LexA) have been shown or are suspected of contributing to control of luminescence (Ulitzur and Dunlap, 1995). Those factors that can restrict growth rate while enhancing expression and activity of luciferase, e.g., limiting oxygen, limiting iron and high or

CHAPTER 1.27

low osmolarity (Dunlap, 1991a; Hastings et al., 1987; Meighen and Dunlap, 1993) might operate by influencing the cellular levels of cAMP and CRP or other stress-response elements.

The Lux Genes, Luminescence Autoinduction and Quorum Sensing The lux Genes. The bacterial lux genes can be grouped in three categories: the core lux, accessory lux and regulatory genes. The five core lux genes, which provide the enzymatic capability for light production, are common to all luminous bacteria examined to date. These genes are luxA and luxB, encoding the luciferase subunits, and luxC, luxD and luxE, encoding the fatty-acid reductase subunits; they occur contiguously as an operon, luxCDABE (Fig. 3). The bacterialluxgenes have been used for a variety of applications, primarily as reporters for environmental and regulatory effects in heterologous systems (LaRossa, 1998). Accessory lux genes, which are associated with light production, are found in different species and strains of luminous bacteria. In some cases, these genes are linked to the lux operon. Photobacterium phosphoreum (Fig. 3) and a strain of P. leiognathi bear luxF, a gene similar in sequence to luxB, between luxB and luxE, encoding a nonfluorescent flavoprotein. The lux operons of the marine luminous bacteria also contain luxG, which in V. fischeri is followed by a strong transcriptional terminator (Swartzman et al., 1990a). The LuxG protein may be a flavin reductase of the Fre/LuxG family of NAD(P)Hflavin oxidoreductases (Zenno and Saigo, 1994a; Zenno et al., 1994b). The last gene of the lux operon in V. harveyi is luxH. Protein LuxH is homologous to E. coli RibB (3,4-dihydroxy-2butanone 4-phosphate synthase), a key enzyme in riboflavin synthesis (Swartzman et al., 1990b). Recently, the ribB gene of V. fischeri was identified. In contrast to luxH in V. harveyi, ribB in V. fischeri is unlinked to the lux operon; nonetheless its expression is controlled coordinately with the lux operon (Callahan and Dunlap, 2000). Additional genes involved in riboflavin synthesis have been identified downstream of the lux operon in V. fischeri, P. leiognathi and P. phosphoreum (Lee et al., 1994c). Other accessory luminescence genes have been described (O’Kane and Prasher, 1992; Meighen, 1994). The third category, genes specifying regulatory proteins, has been identified to date only in V. fischeri and V. harveyi (Fig. 3). In V. fischeri, the main lux regulatory genes, luxR, encoding the acyl-homoserine lactone receptor/lux operon transcriptional activator, and luxI, encoding acylhomoserine lactone synthase (Engebrecht et al., 1983; Engebrecht and Silverman, 1984; Schaefer

CHAPTER 1.27

luxC

luxD

luxA

luxB

luxE

V

luxC

luxD

luxA

luxB

luxE

Pp

luxC

luxD

luxA

luxB

luxF

luxC

luxD

luxA

luxB

luxE

luxC

luxD

luxA

luxB

luxE

Vf

luxR

Luminous Bacteria

luxI

ERIC

875



luxE

ERIC

Fig. 3. Organization of the luminescence genes in luminous bacteria. Abbreviations and key references are: Vf, V. fischeri (Baldwin et al., 1989; Callahan and Dunlap, 2000; Devine et al., 1988a; Engebrecht et al., 1983; Engebrecht and Silverman, 1984; Engebrecht and Silverman, 1987; Foran and Brown, 1988; Swartzman et al., 1990a); Vh, V. harveyi (Cohn et al., 1985; Johnston et al., 1986; Johnston et al., 1989; Miyamoto et al., 1988a; Miyamoto et al., 1988b; Miyamoto et al., 1989; Swartzman et al., 1990b); Pp, P. phosphoreum (Soly et al., 1988); Pl, P. leiognathi (Baldwin et al., 1989; DeLong et al., 1987; Illarionov et al., 1990; Meighen, 1991; Meighen and Dunlap, 1993); Phl, Ph. luminescens (Frackman et al., 1990; Frackman et al., 1990; Johnston et al., 1990; Szittner and Meighen, 1990; Xi et al., 1991); and ERIC, enteric repetitive intergenic consensus sequence (Meighen and Szittner, 1992; Forst and Nealson, 1996). Additional regulatory genes in V. fischeri and regulatory genes in V. harveyi have been identified, as shown in Figs. 4 and 5. Regulatory genes controlling luxCDAB(F)EG expression in other species have not yet been identified. Arrows indicate direction of transcription, and genes are not drawn to scale.

et al., 1996; Stevens et al., 1994; Stevens and Greenberg, 1997), are contiguous with the lux operon. The luxI gene is part of the lux operon, whereas luxR is upstream and divergently expressed (Fig. 3). In V. harveyi, the luxR gene, which is not homologous to V. fischeri luxR, is not linked to the lux operon (Showalter et al., 1990; Swartzman et al., 1992). In other species, the lux regulatory genes have not been identified, but they apparently are unlinked to the lux operon. Additional lux regulatory genes have been identified in V. fischeri and V. harveyi and are described below. Luminescence Autoinduction and QuorumSensing. Many luminous bacteria exhibit a distinctive pattern of luciferase synthesis and light production in laboratory culture, previously called “autoinduction” and now referred to as “quorum sensing.” In V. fischeri and V. harveyi, expression of the lux operon, i.e., luciferase synthesis and luminescence, initially low in early exponential phase cultures, induces strongly as cultures attain the high cell densities associated with late exponential to early stationary phases of growth (Hastings and Greenberg, 1999; Dunlap, 2000). Early analyses of the “phases of luminescence” in culture (Baylor, 1949; Farghaly, 1950) were followed by the demonstration that

luciferase synthesis is inducible and that complete medium contained a compound inhibitory to induction (Nealson et al., 1970; Eberhard, 1972). During growth, cells of V. fischeri and V. harveyi were found to release into the medium species-specific secondary metabolites, called “autoinducers.” These compounds accumulate in the growth medium in a cell-density dependent manner, and once they attain threshold concentrations they induce luciferase synthesis (Nealson et al., 1970; Eberhard, 1972; Nealson, 1977a; Nealson and Hastings, 1979b; Ulitzur and Hastings, 1979; Rosson and Nealson, 1981). The cell density-dependent nature of autoinduction led to the coining of the term quorum sensing (Fuqua et al., 1996; Greenberg, 1997). Analysis of autoinduction reached a notable fruition in the 1980s with the identification of autoinducer signal molecules and lux regulatory genes. The first autoinducer, 3-oxo-hexanoylHSL (3-oxo-C6-HSL), and the first lux regulatory genes, luxI (encoding 3-oxo-C6-HSL synthase; Schaefer et al., 1996) and luxR (encoding acyl-HSL receptor/transcriptional activator) were identified in V. fischeri (Eberhard et al., 1981; Engebrecht et al., 1983; Engebrecht and Silverman, 1984), followed by identification of 3-hydroxybutanoyl-HSL (3-OH-C4-HSL) and a

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nonhomologous luxR gene in V. harveyi (Cao and Meighen, 1989; Showalter et al., 1990). From that base of knowledge, quorum sensing systems that are chemically and genetically homologous to those of V. fischeri and V. harveyi have been identified over the past decade in many species of nonluminous Gram-negative bacteria, including many terrestrial species and several pathogens of animals and plants. Quorum sensing plays important roles in the biology of these bacteria by regulating a wide variety of different kinds of genes, including those for the production of extracellular enzymes, plasmid transfer, antibiotic synthesis and biofilm formation, as well as luminescence (Fuqua et al., 1996; Dunlap, 1997; Swift et al., 1999). Thus, quorum sensing not only is common to luminous bacteria, but also widespread and evolutionarily conserved among Gram-negative bacteria. Despite the importance of quorum sensing as a regulatory paradigm in Gram-negative luminous bacteria, it should be pointed out that apparently not every luminous bacterium can autoinduce luminescence. Certain strains identified as P. leiognathi lack the lag in luminescence and luciferase synthesis in batch culture that is characteristic of autoinduction (Katznelson and Ultizur, 1977); it is possible that expression of the lux system in these strains is independent of acylHSLs and that luciferase synthesis is essentially constitutive. These considerations highlight the likelihood that luminescence and quorum sensing had separate evolutionary origins, discussed below. QUORUM SENSING CONTROL OF LUMINESCENCE IN V. FISCHERI AND V. HARVEYI. Intensive study of V. fischeri and V. harveyi over the past 20 years has developed the luminescence systems in these two species as prototypes for quorum sensing in bacteria (Bassler, 1999; Dunlap, 1997; Dunlap, 2000; Fuqua et al., 1994; Greenberg, 1997; Hastings and Greenberg, 1999). New and fundamental information, nonetheless, continues to accumulate on how these bacteria control light production and use quorum sensing. Indeed, the simple view that “quorum sensing signals accumulate in a cell density-dependent manner and trigger transcription of genes for the luminescence enzymes,” though still entirely valid, has been replaced. The newer view is that quorum-sensing control of luminescence in these two species is remarkably complex, and that the mechanisms (Figs. 4 and 5) have intriguing genetic homologies and disparities. In V. fischeri, luminescence is controlled by two quorum-sensing signals that coordinate a complex regulatory circuitry (Fig. 4). Major components of the quorum-sensing mechanism are: LuxI, the 3-oxo-C6-HSL synthase; LuxR, the

CHAPTER 1.27

transcriptional regulatory protein, which requires 3-oxo-C6-HSL for activity; GroEL, which is necessary for production of active LuxR; and AinS, octanoyl-HSL (C8-HSL) synthase (Engebrecht et al., 1983; Engebrecht and Silverman, 1984; Schaefer et al., 1996; Adar et al., 1992; Dolan and Greenberg, 1992; Adar and Ulitzur, 1993; Hanzelka et al., 1999; Kuo et al., 1994; Gilson et al., 1995; Kuo et al., 1996). A cell density-dependent accumulation of 3-oxo-C6HSL, a membrane-permeant compound (Kaplan and Greenberg, 1985), triggers induction of luxoperon expression by binding to LuxR, apparently a membrane-associated protein (Kolibachuk and Greenberg, 1993), forming a complex that facilitates the association of RNA polymerase with the lux operon promoter (Stevens and Greenberg, 1997). This activation initiates a positive feedback loop for synthesis of 3-oxo-C6-HSL (e.g., Eberhard et al., 1991), and LuxR/3-oxo-C6-HSL negatively autoregulate luxR expression (Dunlap and Greenberg, 1988; Dunlap and Ray, 1989b). The C8-HSL, which apparently interferes with 3-oxo-C6-HSL binding to LuxR, operates to limit premature lux operon induction (Eberhard et al., 1986; Kuo et al., 1996). Expression of luxR is activated by both cAMP and CRP (Dunlap and Greenberg, 1985b; Dunlap and Greenberg, 1988; Dunlap, 1989a; Dunlap and Kuo, 1992c), which also have other regulatory effects (Shadel and Baldwin, 1991) and thereby provide overall control over quorum sensing. Under anaerobic conditions, which are permissive of luciferase synthesis (Eberhard et al., 1979), a regulator of fumarate and nitrate reduction (Fnr) contributes to lux operon expression (Mü ller-Bretkreutz and Winkler, 1993). Recently, a homolog of the V. harveyi luxO gene was identified in V. fischeri. As is the case in V. harveyi, LuxO in V. fischeri apparently functions as a repressor of luminescence (Miyamoto et al., 2000). For details ofluxgene regulation in V. fischeri, see Dunlap (2000). Luminescence regulation in V. harveyi has several features in common with V. fischeri. Like V. fischeri, V. harveyi uses two different quorumsensing signals, 3-OH-C4-HSL and Vh AI-2, an unidentified compound; two different genes (luxLM and luxS) direct synthesis of these respective signals. Luminescence in both species requires cAMP-CRP, and in both it is dependent on a transcriptional activator protein LuxR, although these proteins are not homologous (Bassler et al., 1993; Bassler et al., 1994b; Chen et al., 1985; Eberhard, 1972; Martin et al., 1989; Nealson et al., 1970; Nealson et al., 1972; Miyamoto et al., 1988b; Miyamoto et al., 1990; Miyamoto et al., 1994; Showalter et al., 1990; Swartzman et al., 1990a; Swartzman and Meighen, 1993; Ulitzur and Yashphe, 1975). Fur-

CHAPTER 1.27

Luminous Bacteria cyaA

877

?

AinR AinS

CRP

C8-HSL

cAMP Fnr

CRP

LexA

LuxR

LuxO

+?

+ +

LuxR

GroEL

+

?

luminescence

LuxR

LuxI 3-oxo-C6-HSL

?

QSR 7

q  q 

AcfA

+

+

+

+

 q

Qsr V

host colonization?

DHBP synthase

riboflavin

QsrP

host colonization

Fig. 4. Model for quorum-sensing control of luminescence and other genes of the quorum-sensing regulon of V. fischeri. Depicted in the upper portion of the figure are regulatory genes, proteins, effectors and positive and negative regulatory circuitry controlling luxR/luxICDABEG expression (summarized from Ulitzur and Dunlap, 1995; Dunlap, 2000; see also Lin et al., 2000). Abbreviations: AdCyc, adenylate cyclase; 3-oxo-C6-HSL, 3-oxo-hexanoyl-homoserine lactone; C8-HSL, octanoyl-homoserine lactone. Key elements of lux regulation are described in the text. The weak activation of lux operon expression by C8-HSL/LuxR is depicted as repression, due to the apparent competitive inhibition by C8-HSL of the interaction between 3-oxo-C6-HSL and LuxR (Kuo et al., 1996). The lower portion of the figure indicates genes downstream of the lux operon that are coordinately controlled with the lux operon, positively by LuxR/3-oxo-C6-HSL and negatively by LuxR/C8-HSL (Callahan and Dunlap, 2000).

thermore, the lux operons of these bacteria are similar in structure (Miyamoto et al., 1988a; Swartzman et al., 1990b), although regulatory genes apparently are not present in the region immediately upstream of luxC (Miyamoto et al., 1988b; Fig. 3). The recent identification of ribB, the V. fischeri homolog of V. harveyi luxH, which like the lux operon genes is controlled by LuxR and acyl-HSLs (Callahan and Dunlap, 2000) further demonstrates the overall similarity of lux genes in these two species. A striking counterpoint to the general similarities in the lux operons and physiological control of luminescence in these bacteria is the qualitative difference in quorum sensing in V. harveyi. Expression of the lux operon in V. harveyi is regulated by a quorum-sensing phosphorelay signal transduction mechanism. The mechanism involves two separate two-component phosphorelay paths, each involving a transmembrane sensor/kinase, LuxN and LuxQ, responsive to a separate quorum-sensing signal (Fig. 5). The luxLM genes are necessary for synthesis of the 3-OH-C4-HSL signal. In the absence of 3-OHC4-HSL, LuxN operates as a kinase, phosphorylating LuxU, a signal integrator, which in turn passes the phosphate on to LuxO; the phosphorylated LuxO represses the lux operon. In the presence of 3-OH-C4-HSL, the activity of LuxN is shifted from kinase to phosphatase, which

draws phosphate from LuxU and thereby from LuxO; the dephosphorylated LuxO no longer represses lux operon expression. A similar activity is carried out by a second, as yet unidentified signal (Vh AI-2), which requires LuxS for its production. Operating via LuxP, a putative periplasmic protein, Vh AI-2 mediates the kinase/ phosphatase activity of LuxQ, which in turn, like LuxN, feeds phosphate to or draws it from LuxO. Previously thought to directly repress lux operon expression, LuxO may operate indirectly, by controlling a negative regulator of luminescence. Expression of luxO itself is subject to repression by LuxT (Bassler, 1999; Bassler et al., 1994a; Cao and Meighen, 1989; Freeman and Bassler, 1999a; Freeman and Bassler, 1999b; Lilley and Bassler, 2000; Lin et al., 2000; Surete et al., 1999b). In a manner possibly analogous to LuxR in V. fischeri, LuxR in V. harveyi is autoregulatory and responsive to 3-OH-C4-HSL (Cao et al., 1995; Chatterjee et al., 1996; Miyamoto et al., 1996). The phosphorelay signal transduction mechanism of V. harveyi appears to differ substantially from the V. fischeri quorum-sensing paradigm of direct acyl-HSL/receptor protein activation of lux operon expression. Despite this difference, evidence is growing that the quorum-sensing systems of these two species have significant overlaps at the genetic level. The first indication of genetic overlap was

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CHAPTER 1.27

LuxR

cAMP CRP

LuxR

+

+

Luminescence

LuxT

LuxO

LuxO

D2

P

LuxLM

LuxS

LuxU H2

3-OH-C4-HSL

Vh AI-2 P

P

H1

P P

D1

LuxN

D1

H1

LuxP

LuxQ

Fig. 5. Model for quorum-sensing control of luminescence in V. harveyi. Regulatory inputs positively (cAMP-CRP and LuxR) and negatively (LuxO) controlling the lux operon and negatively controlling luxO are shown. The circled P indicates the intramolecular phosphate transfer and the separate phosphorylation/dephosphorylation signal transduction circuits mediated by the quorum sensing signals 3-OH-C4-HSL and Vh AI-2. H1 and D1 refer to the conserved histidine and aspartate residues of the sensor kinase and response regulator domains, respectively, of LuxN and LuxQ. H2 refers to a histidine residue of LuxU, which integrates signals from LuxN and LuxQ, and D2 refers to an aspartate residue of LuxO, to which the signal is then transduced. The phosphoryl flow is from H1 to D1 of LuxN, and from H1 to D1 of LuxQ, to H2 of LuxU to D2 of LuxO. In the absence of the quorumsensing signals, LuxN and LuxQ, the depicted phosphoryl flow leads to phosphorylation of LuxO, which may directly or indirectly repress the lux operon. Phosphorylase activity of LuxN and LuxQ is activated by the accumulation of the quorum-sensing signals, leading to dephosphorylation and inactivation of LuxO. Not shown are the membranepermeant nature of the quorum-sensing signals and the membrane association of LuxN, LuxQ, and LuxP. Modified from Bassler (1999).

the finding that the C-terminal half of the V. fischeri AinS protein is 34% identical to the V. harveyi LuxM protein, and the N-terminal portion of V. fischeri AinR (encoded by ainR, a gene downstream of ainS) is 38% identical to the Nterminal portion of V. harveyi LuxN (Gilson et al., 1995). Whether AinR itself, possibly with C8HSL, plays a role in lux regulation (Gilson et al., 1995; Kuo et al., 1994) has not been established. The recent identification in V. fischeri of luxO is another example. The deduced amino acid residue sequence of V. fischeri LuxO is approximately 70% identical to that of V. harveyi (Miyamoto et al., 2000). Furthermore, a gene immediately downstream of luxO in V. fischeri is likely to be a homolog of V. harveyi luxU. These homologies suggest that V. fischeri (like V. harveyi) uses a phosphorelay system for quorum sensing. Consequently, the extent of actual differences between the quorum-sensing mechanisms in V. fischeri and V. harveyi is not yet clear.

Nonetheless, the qualitative difference in the quorum sensing systems of V. fischeri and V. harveyi is surprising. One could reasonably anticipate that the mechanisms regulating light production in these two species would be very similar because these bacteria are closely related evolutionarily, and are metabolically and physiologically very similar. Furthermore, whereas V. fischeri and V. harveyi have ecological differences, the differences do not seem to provide a compelling rationale for the qualitative difference inluxregulation. Vibrio fischeri, a more temperate-water species, colonizes light organs of monocentrid fish and certain sepiolid squids, whereas V. harveyi, a species more abundant in warmer waters and able to utilize more sole carbon and energy sources for growth in laboratory culture, has not been found as a light organ symbiont. However, both species can be isolated from seawater, sediments, gut tracts of marine animals, and from infected crustaceans—habitats in which they commonly are found together (Baumann and Baumann, 1981). This ecological commonality suggests that the physiological and ecological importance of quorum sensing would be very similar in the two species. The mechanistic differences outlined above, however, even with genetic overlaps, indicate that the two systems are substantially different and that the difference might result from subtle ecological differences. An alternative possibility, however, is that the lux operons were acquired separately by V. fischeri and V. harveyi, under different circumstances. For example, selective pressures and chromosomal locations for lux operons may have differed at the times of acquisition, accounting for the different qualities of the regulatory mechanisms. Lateral transfer of the lux operon is discussed below. Regardless, the presence of multiple cross-acting quorum-sensing systems in V. fischeri and V. harveyi most likely indicates the importance in both species of sensing and responding to complex and changing conditions in a variety of different habitats. Quorum-sensing Regulated Genes “Downstream” of lux. An emerging area in luminous bacteria biology is the identification of non-Lux activities controlled by quorum sensing. Quorum sensing is known to control various activities in nonluminous bacteria (Dunlap, 1997; Swift et al., 1999), but studies of quorum sensing in luminous bacteria have focused exclusively on luminescence until very recently. Studies of V. harveyi led to the first demonstration of quorum-sensingregulated activities other than luminescence in luminous bacteria. In V. harveyi, the production of the fatty acid storage product poly-bhydroxybutyrate is controlled in a cell densitydependent manner by 3-OH-C4-HSL (Sun et al., 1994; Miyamoto et al., 1998). The use of acyl-

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HSL inhibitors has revealed that production of exotoxins by V. harveyi is linked to quorum-sensing control of luminescence (Harris and Owens, 1999a; Harris et al., 1999b). Furthermore, LuxO controls not only luminescence but also cell morphology and siderophore production in V. harveyi (Lilley and Bassler, 2000). Consistent with these observations, homologs of V. harveyi LuxO, LuxR and LuxS have been identified in several nonluminous bacteria (Jobling and Holmes, 1997; Klose et al., 1998; McCarter, 1998; Sperandio et al., 1999; Surete et al., 1999b; Joyce et al., 2000; McDougald et al., 2000), indicating that, as is the case for V. fischeri, elements of the V. harveyi quorum-sensing system are widespread in Gram-negative bacteria. In V. fischeri, proteomic analysis of mutants defective in luxR, luxI and ainS recently revealed the presence of several quorum-sensing regulated genes “downstream” of the lux operon (Callahan and Dunlap, 2000; Fig. 4). These genes code for an apparently diverse array of proteins, including proteins contributing to the ability of V. fischeri to colonize its squid host, E. scolopes. The identification of non-lux genes in luminous bacteria controlled by quorum sensing and the characterization of quorum-sensing regulatory genes in nonluminous bacteria serves to demonstrate the generality of quorum sensing and to indicate that quorum sensing and luminescence are functionally separate activities.

Independent Evolutionary Origins of Quorum Sensing and Luminescence Knowledge of quorum sensing developed out of studies of luminescence autoinduction in V. fischeri and V. harveyi during the 1970s and 1980s, as described above. Despite the phenomenological and historical linkage between them, however, luminescence and quorum sensing apparently have separate evolutionary origins. The following considerations lead to this view. 1) The existence of strains that apparently do not regulate luciferase synthesis in an autoinducible manner (Katznelson and Ulitzur, 1977) suggests that bacterial luminescence is not necessarily subject to quorum sensing control. 2) The physical linkage of the lux structural and regulatory genes found in V. fischeri, which if consistently present in other species would imply an evolutionary link between light production and its regulation, is not found in other bacteria, e.g., V. harveyi. Furthermore, other luminous bacteria (e.g., V. harveyi) apparently lack homologs of the V. fischeri LuxR and LuxI proteins, and are very different from either V. fischeri or V. harveyi in the way they regulate light production. For example, luminescence in

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Photorhabdus, which does not exhibit autoinduction (Forst and Nealson, 1996), apparently is controlled at the posttranscriptional level (Wang and Dowds, 1991; Hosseini and Nealson, 1995). 3) Many nonluminous bacteria use acyl-HSLmediated quorum sensing, indicating that quorum sensing is not exclusive to luminous bacteria. 4) Quorum sensing controls several activities in nonluminous bacteria (Dunlap, 1997; Swift et al., 1999) and controls activities in luminous bacteria unrelated to luminescence (Callahan and Dunlap, 2000; Sun et al., 1994; Harris et al., 1999b; Lilley and Bassler, 2000), indicating regulatory significance beyond and unrelated to light production. There appears to be no necessary functional or physical connection between quorum sensing and luminescence. In the absence of that connection, one can then ask why luxCDABE is under quorum sensing control in some species but apparently not in others. We postulate that this apparent discordance points to different origins of lux genes among the different luminous bacteria.

Origin and Lateral Transfer of the lux Genes Evolutionary Origin of Bacterial Luminescence The presence of naturally acquired genes necessary for producing light defines the luminous bacteria. The necessary genes luxA and luxB, encoding the luciferase subunits, and luxC, luxD and luxE, for the fatty-acid reductase subunits, are consistently found together as a cotranscribed unit luxCDABE (Fig. 3). Furthermore, the individual lux proteins have a high degree of sequence identity, 54–88% and 45– 77% for the a- and the b-subunits of luciferases, respectively, and 57–80%, 59–74%, and 59–81% for the fatty acid reductase subunits, LuxC, LuxD and LuxE, respectively (Meighen and Dunlap, 1993). The reason for this conservation as a unit is not known; it might be necessary for efficient light production, perhaps by ensuring an interaction of luciferase and fatty acid reductase that facilitates substrate generation and processing. Conservation as a unit to permit coordinate regulation would not seem to be the reason because quorum sensing coordinately regulates several widely separated sets of genes in V. fischeri (Callahan and Dunlap, 2000). The possibility exists therefore that newly identified luminous bacteria will be found to have luxAB and luxCDE in separate chromosomal locations.

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These considerations lead to questions on the evolutionary origin of bacterial luminescence. Seliger (1987) proposed that bacterial luminescence arose under ecological selection, as a biochemical analog of Darwin’s principle of functional change in structural continuity. A flavoprotein catalyzing fatty acid a-oxidation reactions with low chemiluminescent quantum yields is postulated to have mutated under hypoxic conditions to accept FMNH2 as the flavin cofactor, generating a fortuitously high fluorescence yield, termed “protobioluminescence,” via the 4a-hydroxy-FMNH product. This flavindependent, aldehyde-oxidizing protoluciferase produced sufficient light, and with an appropriate emission spectrum, to be detected by phototactic organisms. Responses to the light by visually cueing animals (e.g., to ingest luminous particles), enhanced the growth of the protobioluminescence emitter by introducing it into the animal’s nutrient-rich digestive system, ensuring the emitter’s survival and presumably leading to selection for more intense light output. It is possible that early evolutionary steps leading to protoluciferase involved oxygen detoxification activity that permitted early anaerobic organisms to survive an increasingly aerobic environment (McElroy and Seliger, 1962; Rees et al., 1998). A single gene has been hypothesized to encode bacterial protoluciferase (O’Kane and Prasher, 1992). Although a single-subunit protoluciferase presumably would have differed somewhat from the modern-day luciferase a-subunit and therefore might have produced light, the inability of either of the modern-day a- or b-subunits alone to produce light in vitro or in vivo (Li et al., 1993) argues against the single-gene hypothesis. Alternatively, bacterial luminescence may have arisen following a gene duplication event postulated to have created luxB from luxA (Baldwin et al., 1979; O’Kane and Prasher, 1992; Meighen and Dunlap, 1993). The association of the fatty-acid reductase genes with luxA might have predated the luxA to luxB gene duplication event. Alternatively, the presence of ERIC sequences flanking luxA and luxB in P. luminescens (Meighen and Szittner, 1992) might mark an insertion of the luxAB genes into the fatty aldehyde reductase operon during the evolution of the bacterial luminescence system. Origins and functions of other luminescence proteins have been discussed elsewhere (O’Kane and Prasher, 1992; Meighen and Dunlap, 1993). A marine origin for bacterial luminescence, though speculative, seems reasonable. Most species of luminous bacteria are marine (Table 1), luminescence appears to have arisen independently in various (mainly marine) phylogenetic groups (Hastings, 1995), and present-day luminous organisms are much more common in the

CHAPTER 1.27

ocean than in terrestrial and freshwater environments. Palmer and Colwell (1991) have interpreted the high level of nucleotide sequence identity for a region of luxA among V. cholerae and marine vibrios as indicating a common luminescent marine ancestor. However, a growing number of terrestrial luminous species are being identified (Fischer-Le Saux et al., 1999), so the possibility of a terrestrial origin for bacterial luminescence should not be ruled out.

Lateral Transfer Despite the conservation of the luxCDABE genes in luminous bacteria, the presence of these genes is not monophyletic. Genera with luminous members include the closely related and physiologically similar Vibrio and Photobacterium and the more distantly related and physiologically distinct Photorhabdus and Shewanella (Fig. 2). Various evolutionary scenarios can be envisioned to account for the polyphyletic distribution ofluxgenes and to accommodate the presence of luminous and nonluminous species and strains in Vibrio and Photobacterium: 1) The lux genes may have been present in the ancestor that diverged into the lines leading to modern-day members of the Vibrionaceae, Enterobacteriaceae and Alteromonadaceae. The lux genes were then lost from many descendents but retained by some. If this scenario is correct, one might expect to find more species in the Enterobacteriaceae and Alteromonadaceae that carry lux genes. 2) Alternatively, the lux genes might have arisen later, within the line leading to modernday members of the Vibrionaceae. These genes then may have been lost from several descendents, retained by some, and transferred relatively recently from a member or members of the Vibrionaceae to Photorhabdus and S. hanedai and S. woodyi. The presence of the luxCDABE genes in Photorhabdus species has been interpreted as an instance of lateral gene transfer (Forst et al., 1997). Furthermore, the chromosomal locations of the luxCDABE genes in two ecologically distinct strains of Ph. luminescens apparently differ (Meighen and Szittner, 1992), raising the possibility that lateral transfer to this species occurred more than once (Forst et al., 1997). 3) Also possible is that the lux genes did not arise indigenously in the ancestral line that diverged into the Vibrionaceae, Enterobacteriaceae and Alteromonadaceae (scenario 1) or later within the Vibrionaceae (scenario 2). Instead, they may have been acquired relatively recently by certain species and strains in the Vibrionaceae by lateral gene transfer from an

CHAPTER 1.27

unknown source. The same source might have transferred lux genes to Photorhabdus, S. hanedai and S. woodyi, or these genes might have been acquired secondarily by lateral transfer from a member or members of the Vibrionaceae. The recent identification of luminous strains of V. salmonicida (Fidopiastis et al., 1999) and P. angustum (K. Kita-Tsukamoto et al., manuscript in preparation), species previously characterized as nonluminous, is consistent with all three scenarios. Mapping the chromosomal locations of the lux genes in Vibrio and Photobacterium would help differentiate among these scenarios. Similar chromosomal locations for the lux genes would tend to support an evolutionary origin in an ancestor of or within the Vibrionaceae lineage (scenarios 1 and 2), whereas different chromosomal locations, as seen in Ph. luminescens (Meighen and Szittner, 1992), would be more consistent with lateral transfer to members of the Vibrionaceae (scenario 3). In regard to this latter possibility, the differences in DNA flanking luxCDABE in different members of the Vibrionaceae, for example in V. fischeri and V. harveyi, are intriguing. An issue that complicates each of these scenarios, however, is the possible mobility of the lux genes among members of the Vibrionaceae, with losses and recent lateral transfer events accounting for or contributing to the modern-day presence of luminous and nonluminous species and strains in this family.

Physiological Functions of the Luminescence System One the most interesting and long-standing questions about luminous bacteria is the physiological function of luminescence. In other words, “Why do bacteria produce light?” Despite extensive knowledge of the biochemistry and genetics of bacterial luminescence, the cellular role of luminescence in bacteria is not well understood. However, the benefit light production provides to bacteria has been variously hypothesized, and multiple ecological and physiological functions for luminescence seem likely. Ecologically, in the realm of visually orienting animals, light production undoubtedly plays a role in the dissemination of luminous bacteria and may have been instrumental in the evolution of strongly luminous strains, as discussed above. The feeding of animals on luminous particles (decaying tissues, fecal pellets, and moribund animals infected by luminous bacteria) disperses the bacteria and brings them into the animal’s nutrient-rich gut tract for further growth and dispersal (Nealson and Hastings, 1992). Bioluminescent symbiosis serves a similar role through a continual or diurnal release from the host ani-

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mal, as mentioned above. Once bacteria developed the ability to emit light at levels that could be detected by animals, ecological interactions may then have selected for higher levels of light production (Seliger, 1987), fostering the development of progressively more specific, luminescence-based associations with animals, eventually leading to species-specific bioluminescent symbiosis. A high level of activity of the luminescence system might promote bacterial survival and growth in these associations, especially in bioluminescent symbiosis (Dunlap, 1984; Visick et al., 2000). A strong case for a physiological role for bacterial luminescence can be made, despite the fact that the luxg enes are not essential for survival or growth of luminous bacteria, at least in laboratory culture (Kuo et al., 1994). Several genes are committed to the light-producing reaction, including structural genes for the luminescence proteins and regulatory genes controlling lux operon expression. Retention of these genes and the expenditure of energy for the synthesis and activity of their protein products (Dunlap and Greenberg, 1991b; Meighen and Dunlap, 1993) imply that the luminescence system carries out an activity beneficial to the cell. For example, luciferase activity can consume up to 20% of the oxygen taken up by luminous cells (Eymers and van Schouwenberg, 1937; Watanabe et al., 1975; Dunlap, 1985a; Makemson, 1986a). Furthermore, luminescence in many species is regulated, such that the lux genes are expressed under certain environmental conditions but not others, and lux regulation is deeply integrated into the physiologial response networks and gene regulatory circuitry (e.g., cAMP-CRP and quorum sensing) of the cell. Indeed, luminescence is just one of a suite of metabolic and physiological activities controlled by quorum sensing (Callahan and Dunlap, 2000). The coordinated expression of the lux genes with other sets of genes in response to the physiological state of the cell suggests that the luminescence system plays an integral physiological role in the biology of luminous bacteria. Most attention to the question of that physiological role for bacterial luminescence has focused on oxygen. McElroy and Seliger (1962) proposed that light-emitting reactions arose evolutionarily as detoxifying reactions that removed oxygen and thereby allowed anaerobic organisms to survive. This hypothesis has been developed further from the perspective that luciferin substrates for luciferase are the evolutionary core of bioluminescent systems (Rees et al., 1998). The luminescence reaction, as a terminal oxidase or secondary respiratory chain that is active when oxygen or iron levels are too low for the cytochrome system to operate, would allow

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reduced coenzymes to be reoxidized, thereby permitting cells under microaerobic conditions, such as in animal gut tracts, to continue to metabolize growth substrates (Nealson and Hastings, 1979b; Ulitzur et al., 1981; Makemson and Hastings, 1982; Seliger, 1987). Consistent with this possibility, luciferase activity can partially complement the lack of cytochromes (Makemson and Hastings, 1986b). In bioluminescent symbiosis, the luciferase reaction has been proposed to help protect cells from host-generated reactive oxygen species (ROS; Visick et al., 2000). Alternatively, the physiologically important function of the luciferase reaction may be the production of FP390 (P-flavin binding protein), including its prosthetic group, Q (P)-flavin (Kasai, 1997), according to the following reaction scheme: luciferase

RCHO + FMN + O 2 æ ææ æÆ P-flavin + H 2O + hv The protein FP390 functions as a substitute for flavodoxin, at high salt concentrations where flavodoxin is less active. Flavodoxin, e.g., FldA from V. fischeri (Kasai, 1999), functions to reactivate oxidatively inactivated cobalamindependent methionine synthase (CDMS; Hoover and Ludwig, 1997). It follows that bacteria would not produce light under conditions of low ionic strength because under these conditions cells would use flavodoxin for reoxidation of CDMS in lieu of producing FP390 (Farghaly, 1950; Kasai, 1997). It is tempting to speculate that the postulated relationship between conditions of high ionic strength, synthesis of P-flavin, and light production might account for the apparently exclusive occurence of luminous bacteria in habitats of relatively high ionic strength, i.e., seawater, brackish water, and tissues of marine, freshwater and terrestrial animals. In each of the above cases, light production is an incidental though ecologically important byproduct of the luciferase reaction, and not its primary physiological function. An alternative to incidental light production is the recent proposal that bacterial luminescence serves as an internal light source for a photoreactivation-like repair of damaged DNA (Czytz et al., 2000). Studies of UV survival of V. harveyi lux mutants and an E. coli lexA mutant carrying the V. harveyi lux genes (Czytz et al., 2000) suggest that damaged DNA would be the “missing photoreceptor” for bacterial luminescence. It is intriguing that the V. fischeri lux operon lux box is similar to the E. coli LexA protein-binding site (Ulitzur and Kuhn, 1988; Devine et al., 1988b; Baldwin et al., 1989; Shadel et al., 1990a) and that various SOS-response-inducing and DNA-intercalating agents stimulate luminescence in bacteria (Weiser et al., 1981; Ulitzur and Dunlap, 1995).

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The strong link between oxidative stress and DNA damage (Hemnani and Parihar, 1998) is consistent with this role. Possibly, then, the luciferase reaction carries out the dual physiological functions of ROS detoxification and photoreactivation-like DNA damage repair. The induction of lux gene expression at high population density and the coordinate stimulation of superoxide dismutase (Colepicolo et al., 1992) suggest that DNA damage and oxidative stress become more significant in bacteria as nutrients are exhausted and growth begins to slow.

Isolation, Cultivation and Identification of Luminous Bacteria Detailed information on the isolation, cultivation and phenotypic characterization of luminous bacteria can be found in Nealson (1978a), Baumann et al. (1984b), Baumann and Baumann (1981), and Farmer and Hickman-Brenner (1992). Methods and information not otherwise referenced here were introduced to the author by K. H. Nealson and E. P. Greenberg during summer courses at the Marine Biological Laboratory at Woods Hole, Massachusetts.

Isolation Light-emitting bacteria can be isolated from most marine habitats, through direct plating of samples or by enrichment. For direct plating, 0.1–0.2 ml of coastal seawater is spread on nutritionally complete agar plates, such as Seawater Complete (SWC) agar (Nealson, 1978a; see below). Open-ocean water contains fewer bacteria, so cells from 10 ml to 1 liter are concentrated by filtration (pore size 0.2–0.45 mM), and then the filter is placed on SWC agar or a similar medium. Sediments and gut tracts contain higher numbers of bacteria and therefore usually are diluted 1,000 fold or more before spreading 0.1 to 0.2 ml. Media prepared with 4% agar (Baumann et al., 1984b) helps limit the spreading of swarming and gliding bacteria. Various crustaceans (e.g., gammarid and caprellid amphipods) are suitable sources for luminous bacteria, as they can become infected with luminous bacteria and develop a strong luminescence before and for several hours after dying. In a dark room, after dark-adapting for 12–15 min), one can pick out the infected, luminous crustaceans from collected seaweed. In a lighted room, the exoskeleton of the animal is punctured to obtain the hemolymph, which is streaked onto a suitable agar medium. The plates can be incubated at ambient or cool temperatures and are observed after 12–24 h for luminous colonies, which are then picked and streaked to obtain pure cultures.

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Enrichments of marine luminous bacteria can be made from fresh fish (such as mackerel and flounder), other marine animals, and previously frozen fish. The entire animal or portions are placed in a tray and half covered with seawater, allowing part of the animal’s body to be submerged and part to be exposed to air. This enrichment is then incubated at cool temperatures and observed daily in a darkened room. Luminous spots develop on the exposed portions of the animal within one to several days, depending on the temperature, and these are picked and steaked onto a suitable agar medium. Use of 4% agar (Baumann et al., 1984b) is recommended to limit the spreading of gliding and swarming bacteria. Picking of luminous spots and luminous colonies is made easier by working in a darkened room with a red lamp on variable control. The intensity the lamp and angle of illumination can be adjusted so that luminous colonies are bluish and nonluminous colonies stand out as orangered. Sterile toothpicks are convenient for picking luminous colonies.

Storage Storing luminous bacteria on agar slants or in agar stabs for more than a week is not recommended inasmuch as dim and dark variants can easily arise with some species and survival can be poor. Similarly, survival under refrigeration is poor for some species. Lyophylization or storage in liquid nitrogen may be an option if appropriate equipment is available (Baumann et al., 1984b). Storage at ultra-low temperature, e.g., -75∞C to -80∞C, in a cryoprotective medium, however, works well for all species examined. An effective cryoprotective medium for luminous bacteria is filter-sterilized Deep Freeze Medium (2X DFM), prepared with 1% w/v yeast extract, 10% dimethyl sulfoxide (DMSO), 10% glycerol and 0.2M K2HPO4/ NaH2PO4 (pH 7.0). E. F. DeLong recommended 2X DFM, originally developed by R. Rodriquez for storing yeast. For permanent storage of luminous bacteria, a dense culture is prepared by growing the strain to be stored for 12–18 h in a complete liquid medium, adding 0.5 ml each of the culture and 2X DFM to cryovials, briefly vortexing to mix, allowing the mixture to stand for 10 min before placing the vial into the ultralow temperature freezer. Commercial containers that allow a slow rate of cooling work well as does quick freezing in an ethanol bath kept in the ultra-low temperature freezer or chilled with dry ice. Cultures of luminous bacteria stored in this manner retain viability apparently indefinitely when the tubes are kept at constant ultralow temperature.

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Cultivation Most complete marine media, whether prepared with artificial or natural seawater to supply appropriate levels of Na+, Ca2+ and Mg2+, support the growth of luminous bacteria from most habitats. Nealson (1978a) listed and compared various formulations for complete and minimal media. A commonly used complete medium is SWC, prepared with natural seawater diluted to 70% or 75% with distilled water to minimize precipitation, 5 g per liter of tryptone or peptone, 3 g per liter of yeast extract and 3 ml per liter of glycerol, and with 1.5 g per liter of agar for solid medium. Traditionally, SWC has been buffered with 50 mM Tris or HEPES, or 1 g per liter of solid calcium carbonate has been incorporated into the agar medium to control acid production (Nealson, 1978a). Acid production in SWC apparently results, however, from the presence of glycerol, and elimination of this component avoids the problem (Dunlap et al., 1995) with no major effect on growth or luminescence. An easily prepared complete medium contains 10 g per liter of tryptone, 5 g per liter of yeast extract, 70% natural or artificial seawater, and 1.5% agar for solid medium. Artificial seawater can be prepared according to the formulation of MacLeod, as described by Nealson (1978a), or for routine culture work, a commercial aquarium marine salt mix can be used. Procedures for preparing minimal media have been described by Nealson (1978a).

Identification A combination of phenotypic and genotypic traits is useful for the identification of luminous bacteria. Taxonomy of the marine luminous bacteria and their relationships to other marine enterobacteria were established during the 1970s and early 1980s through the use of an array of diagnostic physiological and molecular traits (Reichelt and Baumann, 1973; Reichelt et al., 1976; Baumann and Baumann, 1981). Using as few as 10–25 phenotypic traits, one can identify with good accuracy many of the commonly encountered species of marine luminous bacteria (Nealson, 1978a; Baumann and Baumann, 1981; Hastings and Nealson, 1981). Genotypic traits, specificallyluxgenes and 16S rRNA (Haygood, 1990; Haygood et al., 1992; Haygood and Distel, 1993b; Wimpee et al., 1991; Nealson et al., 1993), complement these diagnostic characters and can be particularly useful for rapid identification.

Literature Cited Adar, Y. Y., M. Simaan, and S. Ulitzur. 1992. Formation of the LuxR protein in the Vibrio fischeri lux system is

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marine luminous bacteria by using in vitro DNA amplification. Appl. Environ. Microbiol. 57:1319–1324. Wolfe, C. J., and M. G. Haygood. 1991. Restriction fragment length polymorphism analysis reveals high levels of genetic divergence among the light organ symbionts of flashlight fish. Biol. Bull. 181:135–143. Xi, L., K.-W. Cho, and S.-C. Tu. 1991. Cloning and nucleotide sequences of lux genes and characterization of luciferase of Xenorhabdus luminescens from a human wound. J. Bacteriol. 173:1399–1405. Yamada, K., M. Haygood, and H. Kabasawa. 1979. On fertilization and early development in the pine-cone fish, Monocentris japonicus. Ann. Rep. Keikyo Aburatsubo Marine Park Aquarium 10:31–38. Yang, Y., L. P. Yeh, Y. Cao, L. Baumann, P. Baumann, J. S.-E. Tang, and B. Beaman. 1983. Characterization of marine luminous bacteria isolated off the coast of China and description of Vibrio orientalis sp. nov. Curr. Microbiol. 8:95–100. Yasaki, Y. 1927. Bacteriologic studies on bioluminescence. 1: On the cause of luminescence in the fresh water shrimp,

CHAPTER 1.27 Xiphocaridina compressa (De Haan). J. Infect. Dis. 40:404–407. Yetinson, T., and M. Shilo. 1979. Seasonal and geographic distribution of luminous bacteria in the eastern Mediterranean Sea and the Gulf of Elat. Appl. Environ. Microbiol. 37:1230–1238. Zenno, S., and K. Saigo. 1994a. Identification of the genes encoding NAD(P)H-flavin oxidoreductases that are similar in sequence to Escherichia coli Fre in four species of luminous bacteria: Photobacterium luminescens, Vibrio fischeri, Vibrio harveyi, and Vibrio orientalis. J. Bacteriol. 176:3544–3551. Zenno, S., K. Saigo, H. Kanoh, and S. Inouye. 1994b. Identification of the gene encoding the major NAD(P)H-flavin oxidoreductase of the bioluminescent bacterium Vibrio fischeri ATCC 7744. J. Bacteriol. 176:3536–3543. Zobell, C. E., and H. C. Upham. 1944. A list of marine bacteria including descriptions of sixty new species. Bull. Scripps Inst. Oceanogr. 5:239–292. Zobell, C. E. 1946. Marine Microbiology. Chronica Botanica Company. Waltham, MA.

Prokaryotes (2006) 2:893–955 DOI: 10.1007/0-387-30742-7_28

CHAPTER 1.28 l a i re t caB

sn i xoT

Bacterial Toxins VEGA MASIGNANI, MARIAGRAZIA PIZZA AND RINO RAPPUOLI

Introduction Toxins were the first bacterial virulence factors to be identified and were also the first link between bacteria and cell biology. Cellular microbiology was, in fact, naturally born a long time ago with the study of toxins, and only recently, thanks to the sophisticated new technologies, has it expanded to include the study of many other aspects of the interactions between bacteria and host cells. This chapter covers mostly the molecules that have been classically known as toxins; however, the last section also mentions some recently identified molecules that cause cell intoxication and have many but not all of the properties of classical toxins. Tables 1 and 2 show the known properties of all bacterial toxins described in this chapter, while Figure 1 shows the subunit composition and the spatial organization of toxins whose structures have been solved either by X-ray crystallography or by quick-freeze deep-etch electron microscopy. Abbreviations: SEA–SEI, staphylococcal enterotoxin A through I; TSST-1, toxic shock syndrome toxin 1; SPEA, B and C, streptococcal pyrogenic enterotoxins A, B and C; ETA and B, exfoliative toxins A and B; MHC, major histocompatibility complex; Vb or Vg, T-cell-receptor variable domains; LukF, leucocidin F; PA, protective antigen; RTX, repeats-in-toxin; CryIA, CytB, Gi, Gs, Go, Gt, Golf, GTP-binding proteins; MAPKK1 and 2, mitogen-activated protein kinases 1 and 2; EF2, elongation factor 2; Rho, Rac and Cdc42, GTP-binding proteins that control assembly of actin stress fibers; IL2, 4 and 5, interleukins 2, 4 and 5; TeNT, tetanus neurotoxin; VAMP, vesicle-associated membrane protein; BoNT, botulism neurotoxin; SNAP, synaptosome-associated protein; YOP, Yersinia outermembrane proteins; AvrRxv, plant-pathogen virulence protein; Ipa, invasion-plasmid antigen; ICE, interleukin-converting enzyme; Sop, Salmonella outer-membrane protein; Tir, translational initiation region; CagA, cytotoxinassociated gene A; YpkA, Yersinia protein kinase A. For abbreviations, refer to the footnote in Table 1.

Toxins have a target in most compartments of eukaryotic cells. For simplicity, the toxins are divided into three main categories (Fig. 2): 1) those that exert their powerful toxicity by acting on the surface of eukaryotic cells simply by touching important receptors, by cleaving surface-exposed molecules, or by punching holes in the cell membrane, thus breaking the cell permeability barrier (panel 1); 2) those that have an intracellular target and hence need to cross the cell membrane (these toxins need at least two active domains, one to cross the eukaryotic cell membrane and the other to modify the toxin target) (panel 2); and 3) those that have an intracellular target and are directly delivered by the bacteria into eukaryotic cells (panel 3).

Toxins Acting on the Cell Surface See Tables 1 and 2 for a summary of the principal features of toxins described in this section.

Toxins Acting on the Immune System (Superantigens) Superantigens (Fig. 2, panel 1) are bacterial and viral proteins that share the ability to activate a large fraction of T-lymphocytes. They are bivalent molecules that have been shown to simultaneously bind two distinct molecules, the major histocompatibility complex (MHC) and the Tcell receptor variable domains (Vb or Vg; Kotzin et al., 1993; Fig. 3). Binding of these molecules to MHC class II requires no prior processing and occurs outside the antigen-binding groove. This results in the activation of between 2–15% of all T cells, ultimately leading to T-cell proliferation, the production of a variety of cytokines, and expression of cytotoxic activity. Bacterial superantigens, also known as pyrogenic toxins, comprise a class of secreted proteins mostly produced by Staphylococcus aureus and Streptococcus pyogenes (Bohach et al., 1990; Alouf and Muller-Alouf, 2003). So far, they include the group of staphylococcal enterotoxins (SEA, SEB, SECn, SED, SEE, SEG, SEH and

Immune system (Superantigens)

Toxins acting on the cell surface

Aeromonas hydrophyla Pseudomonas aeruginosa Clostridium histolyticum Bacillus cereus

AhyB

Listeria monocytogenes S. pneumoniae

LLO

Alveolysin

B. alveis

S. pyogenes

SLO

Pneumolysin

C. perfringens

PFO

Nhe

ColH

Aminopeptidase

Bacteroides fragilis

S. aureus

ETA, ETB, and ETD

BFT enterotoxin

S. pyogenes

SPEB

Collagenase, metalloprotease Metalloprotease and collagenase Cell membrane permeabilization Thiol-activated cytolysin, cholesterol binding Thiol-activated cytolysin, cholesterol binding Induction of lymphocyte apoptosis Induction of lymphocyte apoptosis Induction of lymphocyte apoptosis

Elastase, metalloprotease

Metalloprotease, cleavage of E-cadherin Elastase, metalloprotease

Trypsin-like serine proteases

Binding to MHC class II molecules and to Vb or Vg of T cell receptor Binding to MHC class II molecules and to Vb or Vg of T cell receptor Cysteine protease

Mycoplasma arthritidis Yersinia pseudotuberculosis

Binding to MHC class II molecules and to Vb or Vg of T cell receptor

Activity

Staphylococcus aureus and Streptococcus pyogenes

Organism

YPMa

SEA-SEI, TSST-1, SPEA, SPEC, SPEL, SPEM, SSA, and SMEZ MAM

Toxin

-

Chronic inflammation

Complement activation, cytokine production, apoptosis Complement activation, cytokine production, apoptosis

Membrane damage

Transfer of other toxins, cell death

Gas gangrene

Cell death

Collagenolytic activity

Corneal infection, inflammation and ulceration Collagenolytic activity

Hydrolization of casein and elastine

-

-

-

-

+

-

-

-

-

-

ETA, ETB

+

+

Chronic inflammation

Alteration in immunoglobulinbinding properties T-cell proliferation, intraepidermal layer separation Alteration of epithelial permeability

SEB SEC2, SEC3, SED, SEH TSST1, SPEA SPEC

X-ray

T cell activation and cytokines secretion

Consequence

V. Masignani, M. Pizza and R. Rappuoli

Small poreforming toxins

Large poreforming toxins

Cell membrane

Surface molecules

Target

Class of toxin

Table 1. Classes of toxins described in the text, their features and activity.

894 CHAPTER 1.28

Class of toxin

Insecticidal toxins

Other poreforming toxins

Membraneperturbing toxins

RTX toxins

Target

S. aureus

PVL leukocidin (LukS-LukF) g-Hemolysins (HlgAHlgB and HlgCHlgB) b-Toxin

Bacillus thuringiensis

CryIA, CryIIA, CryIIIA, etc

C. septicum

AT

E. coli

A. hydrophila

Aerolysin

HlyE

S. aureus

d-Hemolysin

B. anthracis

P. haemolytica

LktA

PA

A. actinomycetemcomitans

LtxA

E. coli

HlyA A. pleuropneumoniae

B. cereus

CytK

ApxI, ApxII, and ApxIII

B. cereus

Hemolysin II

C. perfringens

S. aureus

S. aureus

a-Toxin

Organism B. anthracis

Toxin

ALO

Activity

Cell membrane permeabilization Cell membrane permeabilization Cell membrane permeabilization Calcium-dependent formation of transmembrane pores Calcium-dependent formation of transmembrane pores Calcium-dependent formation of transmembrane pores Calcium-dependent formation of transmembrane pores Perturbation of the lipid bilayer Perturbation of the lipid bilayer Perturbation of the lipid bilayer Perturbation of the lipid bilayer Perturbation of the lipid bilayer Destruction of the transmembrane potential

Cell membrane permeabilization Cell membrane permeabilization

Induction of lymphocyte apoptosis Binding of erythrocytes

Consequence

Osmotic lysis of cells lining the midgut Osmotic lysis of cells lining the midgut

Cell permeabilization and lysis

Cell permeabilization and lysis

Cell permeabilization and lysis

Cell permeabilization and lysis

X-ray

(Continued)

CryIA, CryIIIA

+

+

+ (model)

+

-

-

Apoptosis Activity specific versus ruminant leukocytes

-

-

-

-

-

HlgB

LukF

+

-

Lysis of erythrocytes and other nucleated cells

Cell permeabilization and lysis

Necrotic enteritis

Hemolytic activity

Necrotic enteritis, neurologic effects

Complement activation, cytokine production, apoptosis Release of cytokines, cell lysis, apoptosis Necrotic enteritis, rapid shock-like syndrome Necrotic enteritis, rapid shock-like syndrome

CHAPTER 1.28 Bacterial Toxins 895

Toxins acting on intracellular targets

Target

Bordetella species

Several species

CDT

Toxin C2 and related proteins

C. difficile

Toxins A and B (TcdA and TcdB) Adenylate cyclase (CyaA) Anthrax edema factor (EF) Anthrax lethal factor (LF) Cytotoxin necrotizing factors 1 and 2 (CNF1, 2) DNT

C. botulinum

E. coli

B. anthracis

B. anthracis

B. pertussis

Bordetella pertussis Vibrio cholerae E. coli C. perfringens

PT CT LT a-Toxin (PLC)

PAETA SHT

Corynebacterium diphtheriae P. aeruginosa S. dysenteriae

B. thuringiensis

BT toxin

DT

B. thuringiensis

Organism

CytA, CytB

Toxin

Transglutaminase, deamidation or polyamination of Rho GTPase DNA damage, formation of actin stress fibers via activation of RhoA ADP-ribosylation of monomeric G actin

ADP-ribosylation of EF-2 N-glycosidase activity on 28S RNA ADP-ribosylation of Gi ADP-ribosylation of Gs ADP-ribosylation of Gs Zinc-phospholipase C, hydrolase Monoglucosylation of Rho, Rac, Cdc42 Binding to calmodulin ATPÆcAMP conversion Binding to calmodulin ATPÆcAMP conversion Cleavage of MAPKK1 and MAPKK2 Deamidation of Rho, Rac and Cdc42

Destruction of the transmembrane potential Destruction of the transmembrane potential ADP-ribosylation of EF-2

Activity

+ + + +

cAMP increase cAMP increase cAMP increase Gas gangrene

Failure in actin polymerization

Cell-cycle arrest, cytotoxicity, apoptosis

Ruffling, stress fiber formation

Ruffling, stress fiber formation.

X-ray

-

-

-

CNF1 (catalytic domain)

+

+

cAMP increase Cell death, apoptosis

-

Breakdown of cellular actin stress fibers cAMP increase

-

+ +

+

-

CytB

Cell death Cell death, apoptosis

Cell death

Osmotic lysis of cells lining the midgut Cytocidal activity on human cells

Consequence

V. Masignani, M. Pizza and R. Rappuoli

Cytoskeleton structure

Signal transduction

Protein synthesis

Table 1. Continued

Class of toxin

896 CHAPTER 1.28

Toxins injected into eukaryotic cells

Class of toxin

Cytoskeleton

Inositol phosphate metabolism

Mediators of apoptosis

Intracellular trafficking

Target

P. aeruginosa C. botulinum S. aureus S. typhimurium S. typhimurium

C3 exotoxin

EDIN-A, B and C SopE

SipA

S. flexneri

IpgD

ExoS

Salmonella species

S. pyogenes Shigella Salmonella Yersinia species

H. pylori

C. botulinum

C. botulinum

C. botulinum

SopB

Vacuolating cytotoxin VacA NAD glycohydrolase IpaB SipB YopP/YopJ

BoNT-B, D, G and F neurotoxins BoNT-A, E neurotoxins BoNT-C neurotoxin

C. tetanii

C. perfringens

Iota toxin and related proteins TeNT

Organism E. coli

Toxin

Lymphostatin

Activity

Rac and Cdc42 activation

ADP-ribosylation of Rho Rac and Cdc42 activation

Cleavage of syntaxin, SNAP-25 Alteration in the endocytic pathway Keratinocyte apoptosis Binding to ICE Cysteine proteases Cysteine protease, blocks MAPK and NFkappaB pathways Inositol phosphate phosphatase, cytoskeleton rearrangements Inositol phosphate phosphatase, cytoskeleton rearrangements ADP-ribosylation of Ras, Rho GTPase ADP-ribosylation of Rho

Block of interleukin production Block of interleukin production Cleavage of VAMP/ synaptobrevin Cleavage of VAMP/ synaptobrevin Cleavage of SNAP-25

Consequence

-

Increased chloride secretion (diarrhea)

Breakdown of cellular actin stress fibers Modification of actin cytoskeleton Membrane ruffling, cytoskeletal reorganization, proinflammatory cytokines production Membrane ruffling, cytoskeletal reorganization, proinflammatory cytokines production

Collapse of cytoskeleton

+

EDIN-B +

+

(Continued)

+ (GAP domain)

-

-

Enhancement of GAS proliferation Apoptosis Apoptosis Apoptosis

-

Vacuole formation, apoptosis

BoNT-A

BoNT-B

-

Increased chloride secretion (diarrhea)

X-ray

+ (Hc domain)

+ (C2I)

-

Flaccid paralysis

Flaccid paralysis

Flaccid paralysis

Spastic paralysis

Chronic diarrhea

Chronic diarrhea

CHAPTER 1.28 Bacterial Toxins 897

Signal transduction

Target

L. monocytogenes

BSH

P. aeruginosa

ExoU

B. cereus

S. typhimurium

SptP

Hemolysin BL (HBL)

H. pylori Yersinia species

CagA YopM

V. cholerae

Yersinia species E. coli EPEC

YopH Tir

Zot

Yersinia species

Shigella flexneri

VirA

YpkA

Yersinia species

YopT

Organism Shigella species Yersinia species

Toxin

IpaA YopE

Activity

Hemolytic, dermonecrotic and vascular permeability activities ?

Tyrosine phosphorylated Interaction with PRK2 and RSK1 kinases Inhibition of the MAP kinase pathway Lysophospholipase A activity ?

Vinculin binding GAP activity towards RhoA, Rac1 or Cdc42 Cysteine protease, cleaves RhoA, Rac, and Cdc42 releasing them from the membrane Inhibition of tubulin polymerization Protein serine/threonine kinase Tyrosine phosphatase Receptor for intimin

Consequence

Increased bacterial survival and intestinal colonization

Modification of intestinal tight junction permeability Food poisoning, fluid accumulation and diarrhea

Enhancement of Salmonella capacity to induce TNF-alpha secretion Lung injury

Inhibition of phagocytosis Actin nucleation and pedestalformation Cortactin dephosphorylation Cytotoxicity

Microtubule destabilization and membrane ruffling Inhibition of phagocytosis

Disruption of actin cytoskeleton

Depolymerization of actin filaments Cytotoxicity, actin depolymerization

-

-

-

-

+

+

+ -

-

-

-

+

X-ray

V. Masignani, M. Pizza and R. Rappuoli

Abbreviations: SEA-SEI, staphylococcal enterotoxins; TSST, toxic shock syndrome toxin; SPE, streptococcal exotoxin; SSA, streptococcal superantigen; SMEZ, streptococcal mitogenic exotoxin z; MAM, Mycoplasma arthritidis mitogen; YPMa, Y. pseudotuberculosis-derived mitogen; ETA and ETB, exfoliative toxins; ColH, collagenase; Nhe, nonhemolytic entertoxin; PFO, perfringolysin O; SLO, streptolysin O; LLO, listeriolysin O; ALO, anthrolisin O; AT, a-toxin; PA, protective antigen; DT, diphtheria toxin; PAETA, Pseudomonas aeruginosa exotoxin A; SHT, Shiga toxin; PT, pertussis toxin; CT, cholera toxin; LT, heat-labile enterotoxin; DNT, dermonecrotic toxin; CDT, cytolethal distending toxin; TeNT, tetanus neurotoxin; RTX, repeats in the structural toxin; Hly, hemolysin; Cry, crystal; BoNT, botulinum neurotoxin; Ipa, invasion plasmid antigen; Sip, Salmonella invasion protein; EDIN, epidermal cell differentiation inhibitor; Sop, Salmonella outer protein; Ipg, invasion plasmid gene; Yop, Yersinia outer protein; GAP, GTPase-activating protein; GAS, group A Streptococcus; Vir, virulence protein; YpkA, Yersinia protein kinase A; Tir, translocated intimin receptor; EPEC, enteropathogenic E. coli; CagA, cytotoxin-associated gene A; SptP, Salmonella protein tyrosine phosphatase; VAMP, vesicle-associated membrane protein; ICE, interleukin-1b-converting enzyme; SNAP, synaptosome-associated protein; MAPKK, mitogen-activated protein kinase kinase; Zot, zonula. occludens toxin; and BSH, bile salt hydrolase.

Toxins with unknown mechanism of action

Class of toxin

Table 1. Continued

898 CHAPTER 1.28

CHAPTER 1.28

Bacterial Toxins

899

Table 2. Toxins classified according to their enzymatic activities. Toxin

Substrate

Effect

Glucosyl-transferases Clostridium difficile toxins A and B

Rho/Ras GTPases

Breakdown of cytoskeletal structure

Deamidases E. coli CNF1 Bordetella DNT

Rho, Rac and CdC42 Rho

Stress fiber formation Stress fiber formation

ADP-ribosyltransferases DT PAETA PT CT E. coli LT Clostridium botulinum C2 P. aeruginosa ExoS Clostridium botulinum C3

Elongation factor EF-2 Elongation factor EF-2 Gi, Go and transducin Gs, Gt and Golf Gs, Gt and Golf Actin Ras Rho

Cell death Cell death cAMP increase cAMP increase cAMP increase Failure in actin polymerization Collapse of cytoskeleton Breakdown of cellular actin stress fibers

N-Glycosidases Shiga toxin

Ribosomal RNA

Stop of protein synthesis

Metalloproteases Bacillus anthracis LF Clostridium tetanii TeNT C. botulinum BoNTs

Macrophages VAMP/synaptobrevin VAMP/synaptobrevin, SNAP-25

Disruption of normal homoeostatic functions Spastic paralysis Flaccid paralysis

Abbreviations: CNF1, cytotoxin necrotizing factor 1; DNT, dermonecrotic factor; DT, diphtheria toxin; PAETA, Pseudomonas aeruginosa exotoxin A; PT, pertussis toxin; CT, cholera toxin; LT, heat-labile enterotoxin; ExoS, exoenzyme S; LF, lethal factor; TeNT, tetanus neurotoxin; BoNT, botulinum neurotoxin; VAMP, vesicle associated membrane protein; and SNAP-25, synaptosome-associated protein of 25kDa.

1

11 SEB cytoplasm

SEB TCR DR1

H2N

T

COOH

R

Hn

R

2

Diphtheria Toxin

12

cleavage site

T

HOOC

R

active protease

NH2

active site

A

S. aureus Exfoliative toxins (ETA, ETB, ETD)

B B B B B

Shiga Toxin

Hc

H

Hn

R T

Pseudomonas Exotoxin A

13

H

21

A

N

S. pyogenes SPE-B

3

B

A

Hc

Botulinum neurotoxin

C

RGD prosegment

L

T

membrane

Staphylococcus aureus and Streptococcus pyogenes superantigens

A

s-s

N C

membrane

20

A

B

A cytoplasm

Vacuolating cytotoxin (VacA)

22

B

ExoS (GAP domain)

Fig. 1. Structural features of bacterial toxins. (Left) Scheme of the primary structure of each toxin. For the A/B toxins, the domain composition is also shown. The A (or S1 in PT) represents the catalytic domain, whereas the B represents the receptorbinding domain. The A subunit is divided into the enzymatically active A1 domain and the A2 linker domain in Shiga toxin, CT, Escherichia coli LTI and LTII, and PT. The B domain has either five subunits, which are identical in Shiga toxin, CT, and E. coli LTI and LTII and different in size and sequences in PT, or two subunits (the translocation [T] and the receptor-binding [R] subunits) in DT, Pseudomonas exotoxin A, botulinum toxin, and tetanus toxin. (Right) Schematic representation of the three-dimensional (3D) organization of each toxin. For Staphylococcus enterotoxin B, the protein is shown in the ternary complex with the human class II histocompatibility complex molecule (DR1) and the T-cell antigen receptor (TCR). For Salmonella SptP, the structure is shown in the transition state complex with the small GTP binding protein Rac1. Similarly, toxin SopE is represented in complex with its substrate Cdc42. In the case of E. coli CNF1 and Pseudomonas ExoS, only one domain has been crystallized. In the case of SipA, a 3D reconstruction of SipA bound to F-actin filaments is also reported. For all toxins, the schematic representation is based on the X-ray structure, except that for VacA, whose structure has been solved by quick-freeze, deep-etch electron microscopy. (Continued)

900

V. Masignani, M. Pizza and R. Rappuoli

4

CHAPTER 1.28 B

14

membrane cytoplasm

Perfringolysin O

15

B

C3 Exotoxin

membrane cytoplasm

Staphylococcus aureus, α-Toxin

24

B B B B B

A

HlgA

S1

Pertussis Toxin

5

6

23

S4 S3 S5 S3 S4

S1

Cdc42 toxin

B

SoPE Salmonella typhimurium

Cholera Toxin, E.coli LTI, LTII

A

16

HlgB

A

B

SipA + actin

25

A

A

B

Clostridium perfringens, α - toxin

LukF-PV, Hlg B

7

LF

17

membrane cytoplasm

SipA Salmonella

26

EF

EF PA

Aerolysin

LF

YopE

27

8 PA membrane cytoplasm

HlyE (E. coli)

1

9

Yersinia YopH

Anthrax lethal and Edema factor

28

18 2

1

2 3 domain

3

CNF1

CrylA

YopM Yersinia pestis

(Rho activating domain)

19

10

L

s-s Hn

L

Nt

Ct Rac1

Hc

H

Hn H

CytB

29

Hc

Tetanus Toxin Hc

Ct

Nt

SptP Salmonella species

Fig. 1. Continued.

SEI), exfoliative toxins (ETA and ETB), the toxic shock syndrome toxin-1 (TSST-1; Dinges et al., 2000), the streptococcal pyrogenic enterotoxins (SPEA and SPEC; Papageorgiou et al., 1999) and streptococcal superantigen SSA (Sundberg and Jardetzky, 1999). These toxins play an important role in diseases such as the staphylococcal toxic shock syndrome induced by TSST-1 (Schlievert et al., 1981), vomiting and diarrhea caused by staphylococcal enterotoxins, and the exanthemas caused by the pyrogenic streptococcal exotoxins. Furthermore, these toxins have been linked to the pathogenesis of several acute or chronic human disease states such as the Kawasaki syndrome (Leung et

al., 1993), which is the leading cause of acquired heart disease among children in the United States, and to the pathogenesis of other lifethreatening events such as food poisoning (Blackman and Woodland, 1995). In addition to their functional similarities, the staphylococcal enterotoxins share a number of genetic and biochemical characteristics, as well as similar primary (Schlievert et al., 1995) and 3D structures (Swaminathan et al., 1992; Prasad et al., 1993; Papageorgiou et al., 1995; Schad et al., 1995). The genes for these toxins are generally carried on plasmids, bacteriophage chromosomes, or other heterologous genetic elements (Lindsay et al., 1998; Zhang et al., 1998), and all

CHAPTER 1.28

Bacterial Toxins

2

1

T

3

A

T

N C

A

901

T

B B

cell membrane receptor endosome

Signal

A

Diphtheria Botulinum Tetanus

Toxic factor

Golgi apparatus

Cholera Shiga LT

endoplasmic reticulum

nucleus Fig. 2. Schematic representation of the three groups of bacterial toxins. Group 1 toxins act either by binding receptors on the cell membrane and sending a signal to the cell or by forming pores in the cell membrane, perturbing the cell permeability barrier. Group 2 toxins are A/B toxins, composed of a binding domain (B subunit) and an enzymatically active effector domain (A subunit). Following receptor binding, the toxins are internalized and located in endosomes, from which the A subunit can be transferred directly to the cytoplasm by using a pH-dependent conformational change or can be transported to the Golgi and the endoplasmic reticulum (ER), from which the A subunit is finally transferred to the cytoplasm. Group 3 toxins are injected directly from the bacterium into the cell by a specialized secretion apparatus (type III or type IV secretion system).

T cell receptor

Vα

Vβ superantigen

peptide

MHC class ll molecule

Fig. 3. Schematic representation of the interaction of a superantigen with a major histocompatibility complex (MHC) class II molecule and T-cell receptor.

of them are translated into a precursor protein containing an amino terminal signal sequence that is cleaved during export from the cell. The mature products are small nonglycosylated polypeptide molecules with molecular weights ranging from 20 kDa to 30 kDa and are moderately stable to chemical inactivation, proteolysis and denaturation by boiling. Staphylococcal and streptococcal superantigens share 20–80% sequence similarity (Fig. 4); in particular, staphylococcal SEA is more related to SEE and SED, whereas SEB has greater homology with SEC, TSST-1, and streptococcal superantigens SPEA and SSA. The overall homology found in the staphylococcal enterotoxins has been suggested to stem from duplication of a gene encoding a common “ancestral” toxin (Iandolo, 1989). Computer analysis of the S. pyogenes genome has revealed the presence of novel superantigen genes, and among them the one coding for the mitogenic exotoxin Z (SMEZ). This toxin is particularly similar to the SPE-C group of superantigens and, although present in all group A streptococci (GAS) strains, it shows extensive

902

V. Masignani, M. Pizza and R. Rappuoli

CHAPTER 1.28

Fig. 4. Multiple sequence alignment of staphylococcal and streptococcal superantigens. Green indicates identity, whereas blue stands for amino acid similarity.

allelic variation. Further genetic characterization has shown that SMEZ is the most potent bacterial superantigen so far discovered and that it strongly contributes to the immunological effects of GAS both in vitro and in vivo by eliciting a robust cytokine production (Unnikrishnan et al., 2002). Three novel streptococcal genes (spe-g, spe-h and spe-j) have been identified from the Streptococcus pyogenes M1 genomic sequence, while a fourth novel gene (smez-2) was isolated from the strain 2035. Of these, SMEZ-2, SPE-G and SPEJ are most closely related to streptococcal

pyrogenic exotoxin SPEC, whereas SPE-H is more similar to the staphylococcal toxins than to any other streptococcal toxin (Proft et al., 1999). Finally, other pyrogenic toxin superantigens recently discovered by genome mining include proteins SPEL and SPEM produced by several isolates of S. pyogenes of the M18 serotype. The corresponding genes are contiguous and coded within a bacteriophage. Both toxins were shown to be lethal in different animal models and to directly participate in the host-pathogen inter-

CHAPTER 1.28

Bacterial Toxins

903

Fig. 5. Comparison of the X-ray structures of SEA (left), SEB (right) and TSST-1 (below). The colors follow the secondary structure succession where the N-terminus is blue, the C-terminus is red, and the long central helix is pale yellow. The zinc atom and the coordination site are colored pink and the cysteines involved in the disulfide bond are dark-red.

action in some acute rheumatic fever (ARF) patients (Proft et al., 2003). Crystallographic structures are currently available for most of the described staphylococcal and streptococcal superantigens, such as SEA (Schad et al., 1995), SEB (Swaminathan et al., 1992), SEC2 (Papageorgiou et al., 1995), SEC3 (Fields et al., 1996), SED (Sundstrom et al., 1996), TSST-1 (Prasad et al., 1993; Prasad et al., 1997), SPEA (Papageorgiou et al., 1999), SPEB (Kagawa et al., 2000), SPEC (Roussel et al., 1997) and SSA (Sundberg et al., 1999). However, primary sequence homology among superantigens does not assure homology in their secondary and tertiary structures, and vice versa; in fact SEA, SEB, SEC and TSST-1, despite their low level of sequence similarity, all fold into very similar 3D structures. Below are the X-ray structures of SEA, SEB and TSST-1 that share a very similar fold despite low levels of sequence similarity that range from less than 20% identity in the case of SEA and TSST-1, to 33% in the case of SEA and SEB. All of these toxins have a characteristic twodomain fold composed of a b-barrel at the N-terminus and a b-grasp at the C-terminus connected by a long a-helix that diagonally spans the center of the molecule (Fig. 5). Moreover, all of these toxins are characterized by a central disulfide bond (with the exception of TSST-1, which has no cysteines) and by a Zn+2 coordination site which is believed to be involved in MHC class II binding (Abrahmsen et al., 1995). The presence of two zinc-binding sites in SpeC indicates different modes in the assembly of the MHC-superantigen-T-cell receptor (TcR) trimolecular complex. The crystal structures of SEB and TSST-1 in complex with an MHC class II molecule, and those of SEC2/SEC3 in complex with a TcR Vb chain have been solved (Li et al., 1998; Fields et

al., 1996). As an example, the complex between SEB and the Vb domain of a TcR is reported (Fig. 6). Superantigen molecules have also been identified in other pathogens, where they represent important virulence determinants. MaM is a T-cell mitogen produced by Mycoplasma arthritidis, which contributes to the acute and chronic inflammatory disease mediated by this organism (Cole and Atkin, 1991). The recently determined X-ray structure of MaM in complex with HLA-DR1 has revealed that this protein has a fold and a mode of binding, which are entirely different from those of the known pyrogenic superantigens (Zhao et al., 2004; Fig. 7). Another superantigenic toxin is the YPMa produced by a subset of Yersinia pseudotuberculosis strains. This 14.5-kDa protein was originally purified from bacterial lysates and found to exert a mitogenic activity on human peripheral blood

Fig. 6. Crystal structure of the complex between SEB (green) and TcR (yellow). The residues involved in hydrogen bonds between the two molecules have side-chains colored in red and blue, respectively.

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CHAPTER 1.28

Fig. 7. Three-dimensional structure of MaM superantigen.

mononuclear cells. Although the precise role of this protein is currently unknown, the data show that YPMa contributes to the virulence of Y. pseudotuberculosis in systemic infection in mice (Carnoy et al., 2000). Other toxins that have long been known as superantigens are the streptococcal pyrogenic exotoxin B (SPEB), a virulence factor with cysteine protease activity produced by all isolates of group A streptococci, and the exfoliative toxins A and B produced by S. aureus (Fig. 2, panels 2 and 3). Although these proteins strongly contribute to the virulence of the corresponding microorganism, their role as mitogenic factors has been disproved when it was shown that all the nonrecombinant forms were in fact contaminated with trace amounts of the SMEZ superantigen (Unnikrishnan et al., 2002). SPEB appears to contribute to S. pyogenes pathogenesis in several ways, including proteolytic cleavage of human fibronectin and vitronectin, two abundant extracellular matrix proteins involved in maintaining host tissue integrity. SPEB causes a cytopathic effect on human endothelial cells and represents a critical virulence factor in human infection and in mouse models of invasive disease. Despite low levels of sequence similarity, this toxin can be considered as a structural homologue of the papain superfamily that also includes the mammalian cathepsins B, K and L (Kagawa et al., 2000). Like other proteases, the enzyme SpeB is produced as an inactive precursor (zymogen) of 40 kDa which, following autolytic cleavage of the N-terminal 118 residues, is converted to the mature, active 27.6-kDa protease. The catalytic site lacks the Asn residue generally present in the catalytic Cys-His-Asn triad, which is in this case substi-

Fig. 8. The three-dimensional structure of the precursor form of streptococcal cysteine protease SpeB. The prosegment (blue) and active protease (yellow-orange) are indicated with different color scales. The solvent-exposed ArgGly-Asp (RGD) motif is violet, and the active site (Cys-47His-195-Trp-212) is buried by the prosegment and is colored in green. The highly conserved finger loop is also indicated (arrow).

tuted by a Trp. The structure also reveals the presence of a surface-exposed integrin-binding Arg-Gly-Asp (RGD) motif that is a feature unique to SpeB among cysteine proteases and is linked to the pathogenesis of the most invasive strains of S. pyogenes (Stockbauer et al., 1999). Sequence analysis performed on more than 200 streptococcal isolates has revealed an overall limited structural variation in SPEB, with the entire active site being completely conserved. Interestingly, the prominent finger loop that extends from the N-terminal domain (Fig. 8) is also invariant, suggesting that antibodies directed against this region could be effective therapeutic agents. The exfoliative toxins ETA and ETB of Staphylococcus aureus are produced during the exponential phase of growth and excreted from colonizing staphylococci before being absorbed into the systemic circulation. They have been recognized as the causative agents in staphylococcal scalded skin syndrome, an illness characterized by specific intraepidermal separation of the layers of skin between the stratum spinosum and the stratum granulosum (Ladhani et al., 1999). The two ETs are about 40% identical, with no apparent sequence homology to other bacterial

CHAPTER 1.28

Fig. 9. Crystal structure of exfoliative toxin A (ETA) of Staphylococcus aureus. The three residues of the catalytic triad responsible for the serine protease activity are colored in magenta.

toxins. Both superantigens have been proved to act as serine proteases, and this enzymatic activity could be one of the mechanisms hypothesized as the cause of epidermal separation. In fact, at least in the case of ETA (Fig. 8), substitution of the active site serine residue with cysteine abolishes its ability to produce the characteristic separation of epidermal layers but not its ability to induce T-cell proliferation (Redpath et al., 1991). The two ETs are about 40% identical, with no apparent sequence homology to other bacterial toxins. The overall structures of ETA and ETB are similar to that of the chymotrypsin-like serine protease family of enzymes, with the catalytic triad being composed of His-57, Asp-102 and Ser-195 (Vath et al., 1997, 1999). Recently, a novel member of the exfoliative group of toxins has been discovered in S. aureus. This protein, termed “ETD,” is encoded within a pathogenicity island, which also contains the genes for a serine protease and the edin-B gene. When injected in neonatal mice as recombinant protein, ETD has been shown to induce exfoliation of the skin with loss of cell-to-cell adhesion in the upper part of the epidermis (Yamaguchi et al., 2002).

Toxins Acting on Surface Molecules Bacteroides fragilis enterotoxin (BFT) is a protein of 186 residues that is secreted into the culture medium. The toxin has a zinc-binding consensus motif (HEXXH), characteristic of metalloproteases and other toxins such as tetanus and botulinum toxins. In vitro, the purified enterotoxin undergoes autodigestion and can cleave a number of substrates including gelatin, actin, tropomyosin and fibrinogen. When added to cells in tissue culture, the toxin cleaves the

Bacterial Toxins

905

33-kDa extracellular portion of E-cadherin, a 120-kDa transmembrane glycoprotein (responsible for calcium-dependent cell-cell adhesion in epithelial cells) that also serves as a receptor for Listeria monocytogenes. In vitro, BFT does not cleave E-cadherin, suggesting that the membrane-embedded form of E-cadherin is necessary for cleavage. BFT causes diarrhea and fluid accumulation in ligated ileal loops. In vitro, it is nonlethal but causes morphological changes such as cell rounding and dissolution of tight clusters of cells. The morphological changes are associated with F-actin redistribution. In polarized cells, BFT is more active from the basolateral side than from the apical side, decreases the monolayer resistance, and causes dissolution of some tight junctions and rounding of some of the epithelial cells, which can separate from the epithelium. In monolayers of enterocytes, BFT increases the internalization of many enteric bacteria such as Salmonella, Proteus, E. coli and Enterococcus but decreases the internalization of L. monocytogenes (Sears, 2001). BFT belongs to a large family of bacterial metalloproteases that usually cleave proteins of the extracellular matrix. Pseudomonas aeruginosa and Aeromonas hydrophila elastases (aminopeptidase and AhyB) and Clostridium histolyticum collagenase (ColH) are the best-known examples (Yoshihara et al., 1994; Cascon et al., 2000; Cahan et al., 2001). Lately, a novel member of this family of protein toxins has been identified in Bacillus cereus. The protein, termed “Nhe” (nonhemolytic enterotoxin), is a 105-kDa metalloprotease, which shares homologies to the abovementioned elastases and collagenases. Biochemical characterization has shown that Nhe possesses both gelatinolytic and collagenolytic activities (Lund and Granum, 1999).

Toxins Acting on the Cell Membrane Protein toxins forming pores in biological membranes occur frequently in Gram-positive and Gram-negative bacteria (Braun and Focareta, 1991). Pore-forming toxins, also known as “lytic factors,” work by punching holes in the plasma membrane of eukaryotic cells, thus breaking the permeability barrier that keeps macromolecules and small solutes selectively within the cells (Sugawara et al., 1997; Gilbert, 2002; Fig. 2, panel 1). Because erythrocytes have often been used to test the activity of these toxins, some of them are also called “hemolysins”; however, whereas erythrocytes appear to be very good targets in vitro, they are never the main physiological targets of this class of proteins in vivo (Tomita et al., 1997).

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The pathogenicity of the toxin-producing organisms in eukaryotes is clearly related to the toxins they produce. Furthermore, pore-forming toxins represent the most potent and versatile tool with which invading microbes damage the host cell (Bhakdi et al., 1994). Cell permeabilization exerted by the toxic activity of these proteins generally results in release of cytokines, activation of intracellular proteases, induction of apoptosis, and finally, death of the eukaryotic cell (Alouf and Geoffrey, 1991). To generate channels and holes in the cell membrane, this class of toxins must be able to fold in a characteristic amphipathic structure typical of porins (Weiss et al., 1991; Cowan et al., 1992), with one side facing the internal hydrophilic cavity, and the other side interacting with the lipid chains or the nonpolar segments of integral membrane proteins. Most of the toxins are produced or stored in a protoxin inactive form. The activation step varies from the cleavage of an Nterm acidic peptide as in the case of melittin, to a Cterm proteolytic cleavage as in aerolysin (van der Goot et al., 1992); in the particular case of the Gram-negative hemolysins (cytolysins), these toxins are usually synthesized as precursor proteins, then covalently modified to an acylated, active form and finally secreted via specific export systems, which differ for various types of hemolysins (Issartel et al., 1991; Stanley et al., 1994). All such steps increase the affinity for the membrane, which appears to be essential for activity. A large proportion of these proteins are produced by Gram-positive bacteria and can be divided into large pore-forming and small poreforming toxins on the basis of the dimension of the holes produced on the plasma membrane and also of the kind of interaction that they establish with the eukaryotic receptor. In addition, the pore-forming, repeats-in-toxin (RTX) family of toxins includes a large group of Ca+2dependent hemolysins (secreted by both Grampositive and Gram-negative bacteria), which are characterized by a conserved glycine- and aspartate-rich motif of nine amino acids (Welch, 1991; Coote, 1992). Given their predominant role on cellular membranes, we have included in this section also the so-called “membrane perturbing toxins” and the insecticidal toxins produced by Bacillus thuringensis.

Large Pore-Forming Toxins This class of cytolysins (Fig. 2, panel 2) comprises more than 20 family members, which are generally secreted by taxonomically diverse species of Gram-positive bacteria and which have the common property of binding selectively to cholesterol on the eukaryotic cell membrane (Alouf

CHAPTER 1.28

and Geoffrey, 1991). Each toxin consists of a single 50- to 80-kDa polypeptide chain, and they are characterized by a pretty remarkable sequence similarity, also suggesting possible similar 3D structures. These proteins are produced by Streptococcus pyogenes, S. pneumoniae, Bacillus, a variety of Clostridia, including Clostridium tetanii and C. perfringens, and Listeria. To date, the best characterized are perfringolysin O (PFO), a virulence factor of Clostridium perfringens, which causes gas gangrene (Rossjohn et al., 1997), streptolysin O, secreted by Streptococcus pyogenes (Kehoe et al., 1987), alveolysin, produced by Bacillus alvei (Geoffroy et al., 1990), and pneumolysin, the major causative agent of streptococcal pneumonia and meningitis (Rossjohn et al., 1998). In addition to its role as a cytolysin, listeriolysin O (LLO), which is an essential virulence factor of Listeria monocytogenes (Gedde et al., 2000) has also been shown to induce lymphocyte apoptosis with rapid kinetics (Carrero et al., 2004). These toxins share a similar mechanism of action, which consists of an interaction of monomeric toxin with target cells via cholesterol (their receptor), followed by oligomerization and insertion into the host cell membrane; this process ultimately results in serious membrane damage with formation of large pores with diameters exceeding 150 Å. All these toxins contain a common motif (boxed in Fig. 10), which is located approximately 40 amino acids from the carboxy terminus; this motif includes a Cys residue, which if oxidized abolishes the toxin’s lytic activities. Lytic activity can be restored only upon addition of reducing agents such as thiols. However, despite their designation as “thiol-activated cytolysins,” thiol activation is clearly not an important property of this group of toxins (Billington et al., 2000). Interestingly, the membranebound receptor, cholesterol, plays an important role in the oligomerization step as well as in membrane insertion and pore formation (Alouf and Geoffrey, 1991). Crystallographic data are available only for the thiol-activated cytolysin (perfringolysin O; PFO; Fig. 1, panel 4) of Clostridium perfringens (Rossjohn et al., 1997). Nevertheless, given the high degree of sequence conservation (Fig. 10) detected within this class of protein toxins (ranging from the 43% identity of PFO and listeriolysin, to the 72% identity of PFO and alveolysin), this structure can be considered the prototype of the entire family (Fig. 11). PFO is an unusually elongated rod-shaped molecule mainly composed of b-sheets; the monomer is made of four discontinuous domains, indicated with different colors in the picture. Domain 1 (green) has an a/b structure

CHAPTER 1.28

Bacterial Toxins

907

Fig. 10. Multiple sequence alignment of proteins belonging to the class of large-pore-forming toxins. Green indicates identity, whereas blue stands for amino acid similarity.

containing a seven-stranded antiparallel b-sheet. Domain 2 (blue) consists mainly of four bstrands, while domain 3 (yellow) is comprised of an a/b/a structure. Finally, domain 4 (red) is folded into a compact b-sandwich consisting of multiple-stranded sheets. The mechanism of membrane insertion is not clear; in fact, no canonical transmembrane domains can be identified along the primary structure and no significant patches of hydrophobic residues can be mapped on the surface of the molecule. Nevertheless, a model of the membrane-bound state, which takes into account the interaction with the cholesterol receptor as the first step for penetration of the hydrophobic bilayer core, has been proposed on the basis of electron microscopy and other experimental data. Several chemical modifications and mutagenesis studies have suggested the cholesterol-

binding site to be located at the tip of domain 4 (Fig. 12), and in particular, it has been mapped within the highly conserved, Trp-rich segment (Michel et al., 1990; Hill et al., 1994). Proteolysis studies have further demonstrated that domain 4 is also the membrane-spanning domain, although the distribution of charged and hydrophobic residues on the b-sheet of this region is not compatible with an insertion into the lipid bilayer. From these studies, it has emerged that only the tip of the b-barrel domain D4 is responsible for membrane insertion and that a major conformational rearrangement takes place during pore formation (Shepard et al., 1998; Shatursky et al., 1999). Taken together, these observations suggest a model of oligomer insertion. After the toxin binds to the cholesterol molecule, the aliphatic side chains neutralize the charged resi-

908

V. Masignani, M. Pizza and R. Rappuoli Domain 1

CHAPTER 1.28 1

2

Domain 2 Domain 3

Domain 4

Fig. 11. Crystallographic structure and domain organization of perfringolysin O (PFO) produced by Clostridium perfringens.

3

4

Fig. 13. General mechanism of assembly for small-poreforming toxins: the stem region is initially folded against in the body of the water-soluble monomer; upon binding to the membranes and oligomerization, it subsequently undergoes conformational rearrangement and promotes insertion into the lipid bilayer.

could lead to and promote the final penetration step. Furthermore, on the basis of recent data, a mechanism has been proposed whereby insertion into the bilayer occurs only after PFO monomers have assembled into a pre-pore state. Monomer-monomer interactions therefore not only promote insertion, but cooperative interactions between PFO monomers appear to be required to drive transmembrane insertion and b-barrel formation (Hotze et al., 2002). Recently, a protein belonging to this class of cytolysins has been identified in Bacillus anthracis and named “anthrolysin O” (ALO). This putative toxin is able to bind erythrocytes and could have a role in the virulence of anthrax (Shannon et al., 2003).

Small Pore-Forming Toxins

Fig. 12. Graphical representation of domain 4 of perfringolysin. The Trp-rich loop along with tryptophan side-chains are colored in green. In blue is the b-sheet probably involved in membrane insertion.

dues present on the b-sheet (blue) of domain 4 and then trigger membrane penetration. Consistent with this model is the hypothesis that the highly hydrophobic Trp-rich loop

The family of small-pore-forming toxins acts by creating very small pores (1–1.5 nm of diameter) in the membrane of host cells, thus allowing their selective permeabilization to solutes with a molecular mass less than 2 kDa. Alpha toxin (a-hemolysin) is the prototype of a group of pore-forming toxins produced by most pathogenic strains of Staphylococcus aureus (Gray and Kehoe, 1984a; Song et al., 1996; Gouaux, 1998; Fig. 1, panel 5); other members of this family include leukotoxins, such as leukocidin F (LukF), leukocidin S (LukS), Panton-Valentine leukocidin (PVL) and g-hemolysin (Prèvost et al., 1995; Tomita and Kamio, 1997; Olson et al., 1999; Pedelacq et al., 1999; Cooney et al., 1993) and the b-toxin of Clostridium perfringens (Steinthorsdottir et al., 2000; Tweten, 2001; Magahama et al., 2003). These staphylococcal and streptococcal proteins are secreted as watersoluble monomers and assemble on the surface of susceptible cells to form heptameric transmembrane channels of approximately 1 nm in diameter (Finck-Barbancon et al., 1993; Sugawara et al., 1997; Fig. 13).

CHAPTER 1.28

Bacterial Toxins

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Fig. 14. Multiple sequence alignment of proteins belonging to the family of small pore-forming toxins. Green and blue stand for amino acid identity and similarity, respectively.

The monomers have molecular weights of 33 kDa and are related in sequence and function (Fig. 14). These toxins bind to human erythrocytes, monocytes, platelets, lymphocytes and endothelial cells, causing (at high concentrations) membrane rupture and cell lysis and death. Alpha-toxin has been recently shown to be the major mediator of caspase activation and apoptosis (Haslinger et al., 2003). The structure of the transmembrane pore of staphylococcal a-toxin has been solved and has

Fig. 15. Top and side views of the heptameric complex of a-toxin; each monomer is represented here with a different color (see Fig. 1, panel 5).

confirmed the heptameric structure of the oligomer (Song et al., 1996; Fig. 15). The complex is mushroom-shaped and measures 100 Å in height and up to 100 Å in diameter; the aqueous channel forms the transmembrane pore and spans the length of the entire complex ranging from 14 Å to 46 Å in diameter. Each protomer (Fig. 16) is mainly composed of b-strand elements; two of these in particular constitute the stem domain, which contributes to the formation of the transmembrane pore in the heptameric form of the complex; a glycine-rich

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CHAPTER 1.28

Stem domain

Fig. 16. Structure of the monomer of S. aureus a-toxin; the stem domain involved in pore formation protrudes outside of the core of the structure.

segment that is probably involved in solvent interaction characterizes this domain. Leukotoxins and g-hemolysin (HgII) should be grouped together, inasmuch as they form two types of bi-component complexes (LukF+LukS and LukF+HgII) that exhibit leukotoxic and hemolytic activity, respectively (Tomita and Kamio, 1997). Panton-Valentine leukocidin (PVL) is a closely related toxin carried by 2% of clinically isolated S. aureus strains and is also composed of type F and S components (Prèvost et al., 1995). The components of each protein class are produced as nonassociated, watersoluble proteins that undergo conformational changes and form oligomeric complexes after recognition of their cell targets, a process leading to transmembrane-pore formation and, ultimately, to cell death. The resultant transmembrane channels (estimated diameter 8 Å) are mainly permeable to divalent cations. Recently, fluorescence microscopy experiments have been performed to elucidate the mechanism of membrane insertion of the g-hemolysin complex. This study shows that the three cooperative stages (dimer-dimer interaction, single pore assembly, and aggregation of pores) enhance the efficiency of assembly of oligomeric pores (Nguyen et al., 2003). As representative of this class of bicomponent toxins, consideration is given the X-ray structure of the Luk-F protomer (Olson et al., 1999; Fig. 17), which has been solved at a 1.90 Å resolution. The superposition of this monomer with that of a-toxin shows that the core structures are very similar despite the relatively low primary sequence identity (32%); nevertheless, a conformational change has affected the region of the glycine-rich stem domain, which appears in this case as a compact b-sheet folded against the body of the structure.

Fig. 17. Crystallographic structure determined for the protomer of toxin LukF; the glycine-rich, stem domain is in this case folded against the main body of the structure.

From a structural point of view, in contrast to a wide range of bacterial and insect toxins that utilize a-helices to perturb or penetrate the bilayer, these pore-forming toxins (members of an emerging family of proteins) can be defined by their use of bilayer-spanning antiparallel b-barrels instead. Since the initial discovery of the first small pore-forming toxins, the number of these proteins has grown to include several members, among which are the recently identified hemolysin II (HlyII), and cytotoxin K (CytK) of Bacillus cereus, implicated in necrotic enteritis (Lund et al., 2000; Hardy et al., 2001; Miles et al., 2002).

RTX Toxins Escherichia coli hemolysin (HlyA) is a 110-kDa protein, which can be considered as the prototype of a class of pore-forming toxins mainly produced by Gram-negative bacterial pathogens (Felmlee et al., 1985; Welch, 1991). This wellrepresented family includes a large number of calcium-dependent cytolysins known as RTX toxins, which are produced by different genera of Enterobacteriaceae and Pasteurellaceae. Characterized by the presence of a conserved repeated glycine- and aspartate-rich motif of nine amino acids, these cytolysins have multiple calcium-binding sites essential for function (Felmlee and Welch, 1988). The toxin is encoded by four genes, one of which, hlyA, encodes the 110-kDa hemolysin.

CHAPTER 1.28

Bacterial Toxins

911

≈ 1 kb

A) C activation

A cytolysin

D B transport

tolC transport

B) C adenylate activation cyclase

A cytolysin

D B transport

E transport

Fig. 18. Schematic representation of the genetic organization of RTX determinants; the genes encoding the Hly, Lkt, Aalt and Hpp proteins are organized in the same fashion, as illustrated in panel A, whereas the genes involved in synthesis and secretion of adenylate cyclase/hemolysin of B. pertussis display a somewhat different organization (panel B).

The other genes are required for its posttranslational modification (hlyC) and secretion (hlyB and hlyD). The four genes are found in a very limited number of E. coli clonal types, and can be sometimes located on transmittable plasmids (Smith and Halls, 1967). To give an idea of the level of toxicity associated with hlyA gene product, when non-hemolytic strains of E. coli are transformed with recombinant plasmids encoding the hemolysin, the transformants (in rodent models of peritonitis) are 10-fold to a 1000-fold more virulent than the parental strains. The receptor-binding domain of HlyA has been recently mapped (Cortajarena et al., 2003). Other members of this class of RTX proteins include the adenylate cyclase/hemolysin of Bordetella pertussis (CyaA; Glaser et al., 1988), the ApxI-II and III hemolysins from Actinobacillus pleuropneumoniae (Maier et al., 1996), and the leukotoxins of A. actinomycetemcomitans (LtxA; Korostoff et al., 1998; Henderson et al., 2003) and of Pasteurella haemolytica (LktA; Chang et al., 1987; Wang et al., 1998). Although a remarkable level of primary structure similarity can be detected among this group of toxins (20–60% identity), nevertheless they differ in host cell specificity and seem to adopt diverse mechanisms for cellular damage (Frey et al., 2002). The synthesis and secretion of RTX toxins involve the participation of at least five different gene products; the organization of the five genes is very similar (Fig. 18, panel A), with the exception of B. pertussis bifunctional adenylate cyclase/hemolysin, where all five (cyaC, A, B, D and E) are found together (Glaser et al., 1988; Barry et al., 1991; Fig. 18, panel B); for the other family members, in fact, four of the genes are encoded within a single operon, whereas the fifth gene is located approximately 1 kb downstream (Welch and Pellett, 1988; Wandersman and Delepelaire, 1990). The activation process performed by HlyC on HlyA ultimately results in the acquired capacity of HlyA to bind target cells; this activation

involves proteolytic processing and posttranslational acylation, as well as binding of Ca+2 ions to the repeated domain.

Membrane-Perturbing Toxins d-Toxin or d-hemolysin is secreted into the medium by S. aureus strains at the end of the exponential phase of growth. It is a 26amino-acid peptide (MAQDIISTIGDLVKWIIDTVNKFTKK) that has the general structure of soap with a nonpolar segment followed by a strongly basic carboxy-terminal peptide. The peptide has no structure in aqueous buffers but acquires an a-helical structure in low-dielectricconstant organic solvents and membranes. The a-helix has a typical amphipathic structure, which is necessary for the toxin to interact with membranes. The toxin binds nonspecifically parallel to the surface of any membrane without forming transmembrane channels. At high concentration, the peptide self-associates and increases the perturbation of the lipid bilayer that eventually breaks into discoidal or micellar structures. Interestingly, mellitin, which is also a 26-amino-acid lytic peptide produced by S. aureus, has no sequence homology with dtoxin but has identical distribution of charged and nonpolar amino acids. These toxins are active in most eukaryotic cells. Cells first become permeable to small solutes and eventually swell and lyse, releasing cell intracellular content. Recent data have demonstrated that d-hemolysin insertion is strongly dependent on the peptide-to-lipid ratio, suggesting that association of a critical number of monomers on the membrane is required for activity. The peptide appears to cross the membrane rapidly and reversibly and cause release of the lipid vesicle contents during this process.

Other Pore-Forming Toxins Additional members of this class of b-barrel, channel-forming toxins include aerolysin of Aer-

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CHAPTER 1.28

Domain 2

Domain 1

Domain 3

Domain 4

Fig. 19. X-ray structure of aerolysin toxin of Aeromonas hydrophila; the four domains are indicated; in particular, domain 1 clearly protrudes outside of the main body of the structure.

omonas hydrophila (Parker et al., 1994; Rossjohn et al., 1998), and the closely related a-toxin of Clostridium septicum (Ballard et al., 1995), the anthrax toxin protective antigen PA of Bacillus anthracis (Petosa et al., 1997; Wesche et al., 1998), and the HlyE pore-forming toxin produced by pathogenic E. coli. AEROLYSIN AND ALPHA-TOXIN (AT). Aerolysin (Fig. 1, panel 7) is mainly responsible for the pathogenicity of Aeromonas hydrophila, a bacterium associated with diarrheal diseases and wound infections (Altwegg and Geiss, 1989; Fivaz et al., 2001). It is secreted as a 52-kDa protoxin that is proteolytically cleaved into a 25residue carboxy-terminal peptide and a 48-kDa active protein. Like other functionally related toxins, aerolysin changes its topology in a multistep process from a completely water-soluble form to a membrane-soluble heptameric transmembrane channel (ca. 1.5 nm in diameter) that destroys sensitive cells by breaking their permeability barriers. Proaerolysin is a dimer in solution as well as in the crystal form (van der Goot et al., 1993; Parker et al., 1994); four structural domains characterize the monomer (Fig. 19). In the structure of the dimer, the position of domain 1 appears to be stabilized by contacts with domain 1 of the other monomer, resulting in a very strict interaction of the two (Fig. 20). Domain 4 is characterized by an amphipatic b-barrel structure, which is responsible for mem-

Fig. 20. Structure of the dimer of aerolysin and interaction between the two first domains.

brane insertion of the final complex. In fact, oligomerization is an essential step in channel formation and it seems to precede membrane insertion. A model has been suggested for the entire process; it assumes that proaerolysin approaches the target cell as a water-soluble, hydrophilic dimer which, once concentrated on the surface of the target cell, binds to the receptor; subsequent proteolytic cleavage would cause dimer dissociation and oligomerization. This would ultimately result in an exposure of the hydrophobic region of the toxin and thus in membrane penetration. Clostridium septicum AT is a channel-forming protein that is an important contributor to the virulence of the organism. Recent data have proved that this toxin, like aerolysin, binds to glycosylphosphatidylinositol (GPI)-anchored protein receptors. Furthermore, AT is also active against Toxoplasma gondii tachyzoites. Toxin treatment causes swelling of the parasite endoplasmic reticulum thus providing the first direct evidence that a-toxin is a vacuolating toxin (Ballard et al., 1995; Gordon et al., 1999). Recently, based on the available crystal structure of aerolysin, a molecular model of the membrane spanning domain of AT has been generated (Melton et al., 2004). ANTHRAX PROTECTIVE ANTIGEN (PA). Anthrax protective antigen (PA; Fig. 1, panel 17) is one of the three components of the anthrax toxin complex secreted by Bacillus anthracis, which also includes the edema factor (EF) and the lethal factor (LF; Brossier et al., 2000; Collier and Young, 2003). Whereas EF and LF are responsible for the toxic activity, PA can be considered as the receptor-binding domain for two distinct A subunits, which are in turn EF and LF. The three subunits are encoded on a plasmid and are synthesized and secreted independently. Once on the host cell surface, PA needs a proteolytic activation to form a membrane-inserting heptamer through which EF and LF can be translocated (Klimpel et al., 1992; Milne and Collier, 1993; Milne et al., 1994; see Fig. 37 for the mechanism of action). The monomer is mainly constituted by antiparallel

CHAPTER 1.28

Bacterial Toxins

913

Domain 4 Domain 3

β-tongue

Domain 1 Domain 2

Fig. 21. X-ray structure of Bacillus anthracis protective antigen PA. The four structural domains are indicated by different colors. The two cysteines present in domain 1 are colored in yellow.

b-sheets and contains four functional domains (Fig. 21). The crystallographic structure has revealed how PA can be assembled into heptamers and has suggested how some of the domains can undergo pH-driven conformational change. Domain 1 (red) contains two Ca+2 ions (yellow) and the cleavage site for proteolytic activation; domain 2 (cyan) is the heptamerization domain and is implicated in membrane insertion; domain 3 (green) has an unknown function, whereas domain 4 (yellow) is for receptorbinding. Given its ability to promote the translocation of many heterologous proteins, PA is being evaluated as a general protein delivery system (Leppla et al., 1999). ESCHERICHIA COLI HLYE. Escherichia coli produces a novel pore-forming toxin HlyE (Fig. 1, panel 8), which is completely unrelated to the E. coli hemolysin HlyA of the RTX family (Reingold et al., 1999; Wallace et al., 2000). Nevertheless, sequence comparison studies confirm the presence of highly homologous toxins in other pathogenic organisms such as Salmonella typhi and Shigella flexneri (these orthologs display 92–98% identity to HlyE). This observation suggests that HlyE could be the prototype of a new family of HlyE-like hemolysins specific for Gram-negative bacteria. This new class of pore-forming toxins form cation-selective water-permeable pores (25–30 Å in diameter); the channel formation could be either part of a mechanism for iron acquisition by the bacterial cell, or it may promote bacterial infection by killing immune cells and causing tissue damage (Ludwig et al., 1999). The crystal structure of HlyE has been solved (Wallace et al., 2000; Fig. 22). The toxin has an elongated shape characterized by a four-helix (A–D) bundle topology with each helix approximately 70–80 Å long. Two pre-

Fig. 22. X-ray structure of E. coli HlyE cytolysin. Two hydrophobic domains are present at the extremities of the a-helical bundle (colored in magenta).

dicted hydrophobic domains have been identified on the primary sequence: both are located at the extremities of the molecule, one being mainly composed of a short b-hairpin (b-tongue) folded between the third and fourth helices of the main bundle, and the other consisting of the C-terminal end (magenta) of helix B (cyan). The precise mechanism of HlyE oligomerization to form the final transmembrane pore is at the moment unknown; nevertheless, the first step involves a process of dimerization of two HlyE molecules that pack in a head-to-tail fashion burying the two hydrophobic patches against each other. Electron microscopy experiments have led to a model of channel formation in which the possible oligomer topology is that of an octameric complex, and the b-tongue domain is primary responsible for interaction with the membrane.

Insecticidal Toxins The class of insecticidal proteins, also known as d-endotoxins, includes a number of toxins produced by species of Bacillus thuringiensis. These exert their toxic activity by making pores in the epithelial cell membrane of the insect midgut (Hofte and Whiteley, 1989; Knowles, 1994). d-Endotoxins form two multigenic families, cry and cyt; members of the cry family are toxic to insects of Lepidoptera, Diptera and Coleoptera orders (Hofmann et al., 1988), whereas members of the cyt family are lethal specifically to the larvae of Dipteran insects (Koni and Ellar, 1994). The insecticidal toxins of the cry family are synthesized by the bacterium as protoxins with molecular masses of 70– 135 kDa; after ingestion by the susceptible insect, the protoxin is cleaved by gut proteases to release the active toxin of 60–70 kDa (Drobniewski and Ellar, 1989). In this form, they bind specifically and with high affinity to protein

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Fig. 23. Comparison of X-ray structures determined for representatives of the cry and cyt families of insecticidal dendotoxins: CryIA (left panel) is organized in three structural domains, whereas CytB (right panel) is a singledomain globular protein.

receptors and create channels 10–20 Å wide in the cell membrane. This subgroup includes several toxins (CryIA, CryIIIA, CryIV, CryV, etc.), whereas the only proteins so far characterized that belong to the cyt are CytA and CytB (Koni and Ellar, 1993; 1994). Three-dimensional (3D) structures determined for members of the two families show that the folding of these toxins is entirely different. As representative of the two families, consideration is given to the structures of CryIA (Fig. 1, panel 9; Grochulski et al., 1995) and CytB (Li et al., 1996; Figs. 1 [panel 10] and 23), which share more than 39% sequence identity, suggesting an overall similar folding of the corresponding 3D structures. The CryIA toxin is a globular protein composed of three distinct (but closely packed) domains connected by single linkers: domain 1 is totally a-helical, domain 2 consists of three antiparallel b-sheets and two short a-helices, and domain 3 is a b-sandwich. On the other hand, CytB (also a globular protein) is composed of a single domain with a/b architecture. The molecular mass of the protoxin is in this case only 30 kDa. The region of CryIA, which has been associated with receptor–binding, maps within a loop of domain 2, whereas domain 1 has been shown to be responsible for membrane insertion and pore formation (Martens et al., 1995); this notion is strongly supported by the high structural similarity between the domain 1 of CryIA and that of CryIIIA to the pore-forming domains of colicin A and diphtheria toxin, both composed of helical bundles (Cabiaux et al., 1997; Duche et al., 1999). Conversely, in the case of the CytB/A, the model that has been proposed for the channel formation is based on a b-barrel structure. Because they are toxic to several species of insects, d-endotoxins have been formulated into

CHAPTER 1.28

commercial insecticides, and these insecticides have been used for more than three decades. Recently, Lepidoptera-specific toxin genes have also been used to engineer insect-resistant plants (Christov et al., 1999). Very recently, a novel crystal protein produced by B. thuringiensis has been identified. This toxin (BT) is noninsecticidal and nonhemolytic, but has strong cytocidal activity against various human cells. Its amino acid sequence has little homology with the other known insecticidal toxins, suggesting that BT might belong to a new group of Bacillus thuringiensis crystal toxins (Ito et al., 2004).

Toxins Acting on Intracellular Targets See Tables 1 and 2 for a summary of the principal features of toxins described in this section. The group of toxins with an intracellular target (A/B toxins) contains many toxins with different structures that have only one general feature in common: they are composed of two domains generally identified as “A” and “B.” The A domain is the active portion of the toxin; it usually has enzymatic activity and can recognize and modify a target molecule within the cytosol of eukaryotic cells. The B domain is usually the carrier for the A subunit; it binds the receptor on the cell surface and facilitates the translocation of A across the cytoplasmic membrane (Fig. 2, panel 2). Depending on their target, these toxins can be divided into different groups that act on protein synthesis, signal transduction, actin polymerization, and vesicle trafficking within eukaryotic cells.

Toxins Acting on Protein Synthesis These toxins are able to cause rapid cell death at extremely low concentrations. Two ADPribosylating bacterial proteins (see also the section ADP-Ribosyltransferases: A Family of Toxins Sharing the Same Enzymatic Activity) are actually known to belong to this class of toxins: diphtheria toxin (DT) of Corynebacterium diphtheriae (Pappenheimer, 1977; Collier et al., 1982) and Pseudomonas aeruginosa exotoxin A (PAETA; Gray et al., 1984b; Wick et al., 1990). Both display their toxic activity by transferring the ADP-ribose moiety to a posttranslationally modified histidine residue of the cytoplasmic elongation factor 2 (EF2) of eukaryotic cells (Brown and Bodley, 1979; Van Ness et al., 1980). This reaction leads to the formation of a completely inactive EF2-ADP-ribose complex, which ultimately results in inhibition of protein

CHAPTER 1.28

synthesis and cell death. From the biochemical point of view, the two toxins have a similar size, a signal peptide and disulfide bridges, and both are produced in iron-depleted medium. Nevertheless, they show a completely different amino acid composition and bind different cell receptors. In addition, Shiga toxin is another protein that exerts its toxic activity by interfering with protein synthesis. DIPHTHERIA TOXIN. This toxin (DT; Fig. 1, panel 11) is a 535-amino acid polypeptide that is secreted into the growth medium by strains of toxinogenic Corynebacterium diphtheriae, and the polypeptide sequence is encoded by a lysogenic bacteriophage. Biosynthesis is regulated by an iron-binding protein, and proceeds only in the absence of iron (Qiu et al., 1995; Ding et al., 1996). The toxin is synthesized as a single polypeptide chain that is subsequently cleaved into two fragments, A and B of 21 kDa and 37 kDa, respectively (Pappenheimer, 1977). From the functional point of view, three separate domains (C, T and R) are seen in the crystallographic structure of DT. The catalytic domain (C) entirely corresponds to the A subunit, whereas the translocation domain (T) and the carboxy-terminal, receptor-binding domain (R) are contained in fragment B (Choe et al., 1992; Bennett and Eisenberg, 1994). From the structural point of view, the C domain (residues 1–191) has an a+b structure, the receptor-binding domain is a flattened b-barrel with a jelly-roll-like topology, whereas the translocation domain T (residues 201–384) consists in nine helices, two of which may participate in the pH-triggered membrane insertion. The molecule contains four cysteines and two disulfide bridges: one joins fragment C to fragment T and the other is contained within fragment R (Fig. 24). Although the toxicity of DT is entirely due to the enzymatic activity carried on by fragment A (Fig. 25), fragment B is absolutely required for the cell intoxication process. After secretion from Corynebacterium diphtheriae, the toxin binds to the DT receptor and is internalized by receptor-mediated endocytosis. In the endosome, the acidic environment triggers a conformational change of the B subunit that exposes the hydrophobic regions of the T domain allowing the interaction with the endosomal membrane and the translocation of the amino-terminal catalytic domain C across the membrane to the cytosol. According to a recent model, the A subunit of DT is able to cross the endosomal membrane making use of a metastable transmembrane domain, which has also been identified (Wolff et al., 2004). The toxin receptor is the heparin-binding, epidermal growth factor (EGF)-like precursor (Naglich et al., 1992;

Bacterial Toxins

915

Fig. 24. X-ray structure of diphtheria toxin. The three functional domains are indicated with different colors: the catalytic domain C is green, the translocation domain T is red and the receptor-binding domain R is cyan. The two disulfide bridges are colored yellow.

His 21

Glu 148 active site loop

Fig. 25. Crystal structure of the isolated catalytic domain of diphtheria toxin. The scaffold of the enzymatic cleft is green, and the two described catalytic residues are blue. The “activesite loop” is represented here in the “closed” conformation.

Hooper and Eidels, 1995) that is present in most mammalian cells; nevertheless, the receptors of murine cells contain a few amino acid substitutions that make rodents insensitive to DT. Diphtheria toxin is one of the most potent bacterial toxins: in vitro experiments have shown that a single molecule of the enzymatically active fragment A is by itself able to kill one eukaryotic cell (Yamaizumi et al., 1978). Biochemical and mutagenesis studies have greatly contributed to the understanding of structure-function relationships and to the mapping of the catalytic residues. In particular,

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His-21 has been mutagenized with a number of different residues and has been found to be essential for catalysis (Papini et al., 1989; Blanke et al., 1994); in fact, some activity was maintained only when Asn replaced His. In a similar manner, Glu-148 was identified as an active-site residue by photoaffinity labeling experiments with nicotinamide adenine dinucleotide (NAD; Carroll et al., 1985) and subsequent site-directed mutagenesis studies; in this case, not even a conservative substitution with Asp could be possible without complete loss of activity (Tweten et al., 1985). Whereas the possible function for His-21 could be that of maintaining the integrity of the activesite pocket, Glu-148 is likely to be involved in the interaction with the upcoming substrate molecule. Later, crystallographic data confirmed and extended the experimental observations, and added a number of other important residues to the list of the catalytic ones. A very important step in the elucidation of the mechanism of enzymatic activity has been the determination of the crystal structure for the complex of diphtheria toxin with NAD (Bell and Eisenberg, 1997). Upon the addition of NAD to nucleotide-free DT crystals, a significant structural change affects the region encompassing residues 39–46. This portion of the C domain constitutes a mobile loop that becomes disordered after the formation of the complex. The best hypothesis to explain this observation is that NAD enters the cavity upon displacement of the mobile loop, which is then made available for the recognition and binding of the acceptor substrate EF-2. This would explain why DT recognizes EF2 only after NAD has bound (see the section A Common Structure of the Catalytic Site in this Chapter). Detoxified diphtheria toxin has been used in the formulation of a vaccine against toxinogenic strains of Corynebacterium diphtheriae (Porro et al., 1980; Rappuoli, 1983). PSEUDOMONAS AERUGINOSA EXOTOXIN A. This exotoxin (PAETA; Fig. 1, panel 12) is a 66-kDa single-chain protein that inhibits protein synthesis (by a mechanism of action identical to that of DT) in eukaryotic cells by catalyzing the transfer of the ADP-ribosyl moiety of oxidized NAD onto elongation factor 2 (Brown and Bodley, 1979; Van Ness et al., 1980; Gray et al., 1984b; Wick et al., 1990; see the section ADP-ribosylating Toxins in this Chapter). Exotoxin A is the most toxic of the proteins secreted by the opportunistic pathogen Pseudomonas aeruginosa, having an LD50 of 0.2 mg upon intraperitoneal injection into mice. Secreted in the supernatant as an enzymatically inactive proenzyme; this toxin must undergo structural alteration to be able to perform its ADP-ribosylating activity.

CHAPTER 1.28

According to X-ray crystallography (Allured et al., 1986; Li et al., 1995; 1996b), the molecule can be divided into three functional domains. The receptor-binding domain I binds to the ubiquitous a2-macroglobulin receptor of eukaryotic cells, thus initiating receptor-mediated endocytosis. This domain is composed primarily of antiparallel b-structure and is arranged in two noncontiguous regions that encompass residues 1–252 (Ia) and 365–399 (Ib), respectively. Domain II maps within amino acids 253–364, is composed mostly of hydrophobic a-helices, and mediates the translocation of the enzymatically active carboxy-terminal domain III (residues 400–613) to the cytosol of infected cells. Furthermore, it has been shown that for domain III to be functional, a specific proteolytic cleavage at residue 280 of domain II is needed. Genetic studies based on the expression of mutated forms of the exotoxin A gene in E. coli have confirmed these functional assignments. In fact, whereas deletion of domain Ia results in nontoxic, enzymatically active molecules that cannot bind the cells, deletions in domain II give rise to molecules that bind to the cells, are enzymatically active, but are not toxic; finally, deletions or mutations in domain III result in enzymatically inactive molecules (Siegall et al., 1989). To become active, the PAETA toxin requires an intracellular furin-mediated proteolytic cleavage to generate a 37-kDa C-terminal fragment that is then translocated to the cytoplasm to reach the EF2 target (Inocencio et al., 1994). By using a fluorescence resonance energy transfer approach, the mechanism of interaction between ExoA and its substrate EF has been studied, showing that the binding is strongly dependent on the pH. Furthermore, the finding that EF-2 bound to GDP or GTP is still recognized by ExoA shows how adaptable this toxin is in ADP-ribosylating its substrate. In particular, mutational analysis affecting the last five residues at the carboxy-terminus of the enzymatic domain resulted in complete loss of cytotoxicity; this segment (Arg-Glu-Asp-LeuLys, REDLK) closely resembles the KDEL motif that is a well-defined endoplasmic reticulum retention sequence and that has also been found at the C-terminus of other ADP-ribosyltransferases such as cholera toxin and heat-labile enterotoxin of E. coli (Chaudhary, 1990). It has been postulated that the sequence REDLK may be a recognition signal required for entry of the ADP-ribosylation domain of PAETA into the cytosol. Four disulfide bonds are present in the structure, but all of them are confined to the portion of exotoxin A that is not required for enzymatic activity. Photoaffinity labeling experiments have identified Glu-553 as an active-site residue; substitu-

CHAPTER 1.28

Bacterial Toxins

917

Fig. 26. Comparison of exotoxin A crystal structures in the absence (left panel) and in the presence (right panel) of the ligand (in red). The active site residues are shown in blue. The loop (when present) is colored in magenta (see Fig. 1, panel 12). Glu 553

tion of this residue with any other amino acid, including the closely related Asp, decreased the enzymatic activity by a factor of 1000 (Douglas and Collier, 1990). In a similar manner, experiments of site-directed mutagenesis on His-440 led to molecules with a severely reduced cytotoxic activity, thus suggesting an important role for this residue in the reaction (Han and Galloway, 1995). The crystal structure of the catalytic domain has been recently solved both in the isolated conformation and in the presence of an NAD analog (b-methylene thiazole-4-carboxamide adenine dinucleotide; b-TAD; Li et al., 1995; 1996b; Fig. 26). Comparison of the two structures shows that the major difference resides in the new conformation of the loop 458–463, which appears to be displaced by ligand binding; displacement of this loop from the active-site cleft could be an essential step allowing entrance and correct positioning of the NAD molecule during the enzymatic reaction. Given the potent lethal activity, the catalytic domain of exotoxin A has been widely used for the construction of fusion proteins with cellbinding domains specific for tumor cells or other types of dangerous cells. So far, nucleotides encoding domain I have been replaced by sequences encoding interleukin (IL) 2, IL-6 and T-cell antigen CD4. These fusion molecules are promising candidates for the treatment of arthritis and allograft rejection (PAETA-IL2), acquired immune deficiency syndrome (PAETACD4), and other diseases (Chaudhary et al., 1987, 1988; Siegall et al., 1988; Ogata et al., 1989; Baldwin et al., 1996; Mori et al., 1997). SHIGA TOXIN. This toxin (SHT; Fig. 1, panel 13), also known as “verotoxin,” is the key virulence factor produced by Shigella dysenteriae, the pathogen responsible for the most severe forms of dysentery in humans (Kozlov et al., 1993). Shiga toxin is the prototype of a family of closely related bacterial protein toxins (Shiga-

His 440

like toxins), also produced by certain strains of E. coli responsible for hemorrhagic colitis (Karmali et al., 1988). From its 3D structure (Fraser et al., 1994), it is possible to recognize this protein as belonging to the class of A/B bacterial toxins, which consist of an enzymatic A subunit associated with a B domain binding to specific cell-surface receptors. The A subunit bears the enzymatic activity and is thus responsible for toxicity; like Pseudomonas aeruginosa exotoxin A and diphtheria toxin of Corynebacterium diphtheriae, SHT has an effect on protein synthesis, and in particular, by means of its N-glycosidase activity, it is able to depurinate a specific adenosine of ribosomal RNA and stop protein synthesis in the target cell (Endo et al., 1988). The catalytic subunit is composed of two regions, A1 and A2, and like many other bacterial protein toxins, it needs to be activated by proteolytic cleavage. Fragment A2 has an a-helical structure and is noncovalently linked to the B domain (Fig. 27). Interestingly, its primary structure displays a notable similarity to chain A of ricin, a plant toxin that also shares the same enzymatic function acting on the same substrate (Katzin et al., 1991). This domain displays an overall organization which is very similar to that of the corresponding receptor-binding subunits of the ADP-ribosyltransferases cholera toxin and heat-labile enterotoxin LT of E. coli, all formed by five identical protomers which assemble into the final ring-like structure of the B oligomer (Fig. 28). The B-subunit of Shiga toxin has been demonstrated as a powerful vector for carrying attached peptides into cells for intracellular transport studies and for medical research (Hagnarelle et al., 2003). Upon binding of verotoxin to its receptor (globotrialosylglyceramide, Gb) on the surface of a eukaryotic cell (Cohen et al., 2000), the toxin is internalized by receptor-mediated endocytosis and is transported to the Golgi and to the endoplasmic reticulum, from which the A subunit is

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CHAPTER 1.28

translocated to the cytoplasm, where it can gain access to the ribosomal target. Numerous recent studies have shown that Shiga toxins trigger programmed cell death signaling cascades in intoxicated cells. The mechanisms of apoptosis induction by these toxins are newly emerging, and the toxins may signal apoptosis in different cells types via different mechanisms (Cherla et al., 2003).

Toxins Acting on Signal Transduction Signal transduction is an essential mechanism for the survival of any living organism. In eukaryotic

cells, signals received from the outside stimulate receptors on the cell surface and are subsequently transmitted across the cell membrane mainly using two types of mechanism: 1) tyrosine phosphorylation of the cytoplasmic portion of the receptor which initiates a cascade of intracellular signaling events; and 2) modification of a receptor-coupled GTP-binding protein that transduces the signal to various enzymes which respond with the release of secondary messengers such as cyclic AMP (cAMP), inositol triphosphate, and diacylglycerol; accumulation of these products alter the normal equilibrium of the cell and thus provoke malfunction and death.

Pertussis Toxin A1

A2

B

Fig. 27. Three-dimensional structure of Shiga holotoxin. The A subunit is distinguished between A1 (blue) and A2 (yellow), whereas the receptor-binding domain B has different colors for the five monomers.

This toxin (PT; Fig. 1, panel 14) is a protein of 105 kDa released into the extracellular medium by Bordetella pertussis, the etiological agent of whooping cough. It belongs to the A/B class of ADP-ribosylating toxins and is composed of five distinct subunits, named “S1” through “S5,” where S4 is present in two copies in the final oligomer. The genes encoding for the five monomers of pertussis toxin are organized into an operon structure (Locht et al., 1986; Fig. 29) and contained within a chromosomic DNA fragment of approximately 3200 base pairs. Interestingly, the genes coding for S2 and S3 share a 75% similarity (67%, if calculated from S2 and S3 gene products at the amino acid level), suggesting a common evolutionary origin for the two sequences, possibly because of gene duplication. The five subunits are independently secreted into the periplasmic space, where the toxin is assembled and then released in the culture

Fig. 28. Bottom view of the B subunit of Shiga toxin (left panel) in comparison to the B subunit of E. coli LT (right panel).

S5 5¢

S4 S1

promotor

S2

terminator S3

3¢

Fig. 29. Schematic representation of the genetic organization of the open reading frames (ORFs) coding for the five subunits of pertussis toxin.

CHAPTER 1.28

Bacterial Toxins

Fig. 30. Three-dimensional structure of the pertussis holotoxin. Left panel: side view of the intact holotoxin; right panel: bottom view of the receptorbinding domain. Each subunit is colored accordingly to the corresponding genes as represented in Fig. 29.

919

S1

S2 - S3 - (S4)2 -S5

medium by a specialized type IV secretion apparatus (Covacci and Rappuoli, 1993a; Weiss et al., 1995). Subunit S1 represents the enzymatically active domain A, which is totally responsible for the toxicity, whereas the pentamer S2-S3-(S4)2S5 constitutes the receptor-binding domain B (Fig. 30). The A domain acts on eukaryotic cells by ADP-ribosylating their GTP-binding proteins, and specifically it transfers an ADP-ribose group to a cysteine residue located in the carboxyterminal region of the a-subunit of many G proteins such as Gi, Go and transducin (Katada et al., 1983; West et al., 1985); Gs which has a tyrosine residue in place of the cysteine is not a valid substrate for PT. The consequence of ADPribosylation is the uncoupling of G-proteins from their receptors which results in an alteration of the response of eukaryotic cells to exogenous stimuli and thus in a variety of in vivo phenotypes, such as leukocytosis, histamine sensitization, and increased insulin production (Sekura, 1985). Conversely, the most interesting activity displayed by PT in vitro is the observed change in cell morphology in Chinese hamster ovary (CHO) cells (Hewlett et al., 1983). The B domain is a nontoxic oligomer that binds the receptors on the surface of eukaryotic cells and allows the toxic subunit S1 to reach its intracellular target proteins through a mechanism of receptor-mediated endocytosis, likely following a mechanism of retrograde transport through the Golgi apparatus. The importance of the Golgi localization of pertussis toxin for the S1-dependent ADP-ribosylation of G-proteins was investigated employing Brefeldin A (BFA) treatment to disrupt Golgi structures. This treatment completely blocked the pertussis toxin ADP-ribosylation activity of cellular G-proteins, therefore indicating that retrograde transport to the Golgi network is a necessary prerequisite for cellular intoxication (el Baya et al., 1997). In CHO cells, the PT receptor has been shown to be a high-molecular weight glycoprotein that binds the B oligomer through a branchedmannose core containing N-acetylglucosamine (Sekura, 1985). In contrast to the other ADP-

ribosyltransferases, where the enzymatically active domain A mediates all the toxic activities, PT possesses other nonlethal activities (such as a mitogenic activity on T cells), which are mediated exclusively by the receptor-binding domain B (Tamura et al., 1983). The active site of pertussis toxin is structurally homologous to the active sites of other ADP-ribosylating toxins. This aspect will be described in the section ADPRibosyltransferases: A Family of Toxins Sharing the Same Enzymatic Activity in this Chapter. Pertussis toxin plays a central role in the pathogenesis of whooping cough and in the development of protective immunity against reinfection. For this reason, the role of many residues of S1 has been tested by site-directed mutagenesis to produce nontoxic mutants of the toxin to be used as vaccines. The minimal region still enzymatically active is constituted by amino acids 4–179 of S1 subunit (Pizza et al., 1988; Cieplak et al., 1988; Fig. 31), and it is within this fragment that many mutations have been

Fig. 31. Crystal structure of the wildtype S1 subunit of pertussis toxin. The scaffold of the enzymatic cleft is represented as a green ribbon, whereas the rest of the molecule is in pale yellow carbon trace representation. Residues proved to be essential for activity by means of site-directed mutagenesis are represented with side chains and are colored in blue.

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CHAPTER 1.28 PT-9K/129G

Lys9

Glu129

Fig. 32. X-ray representation of the wildtype pertussis toxin (left panel) and of the double mutant 9K/129G (right panel). The catalytic cleft is colored in yellow, whereas the mutated residues are in red.

Gly129

designed and analyzed for activity. In particular, Arg-9, Asp-11, Arg-13, Trp-26, His-35, Phe-50, Glu-129 and Tyr-130 were found to be essential for enzymatic activity and, when replaced with other residues, the toxicity was reduced to levels of about 1%; nevertheless, none of the singleamino acid mutants were completely devoid of toxicity. The most successful mutant contains in fact two amino acid substitutions: Arg-9/Lys and Glu129/Gly (PT-9K/129G; Fig. 32). This mutant is structurally identical to the wildtype but is completely nontoxic and has been used for the construction of an acellular vaccine against pertussis. This vaccine has been extensively tested and has been shown to induce protection from disease (Pizza et al., 1989; Rappuoli, 1997).

Cholera Toxin and Heat-Labile Enterotoxin Cholera toxin (CT) and E. coli heat-labile enterotoxins (LT-I and LT-II) share an identical mechanism of action and homologous primary and 3D structures (Dallas and Falkow, 1980; Spicer et al., 1982; Sixma et al., 1991; Figs. 1 [panel 15] and 33). The CT is produced by Vibrio cholerae (the etiological agent of cholera), whereas LT-I and LT-II are produced by enterotoxigenic strains of E. coli (ETEC) isolated from humans with traveler’s diarrhea, from pigs (LTI), or from food (LT-II; Seriwatana et al., 1988). The two toxins belong to the class of ADPribosylating toxins and are organized in an AB5 architecture, where the B domain is a pentamer which binds the receptor on the surface of eukaryotic cells, and domain A bears the enzymatic activity and is thus responsible for toxicity (Holmgren, 1981; Moss and Vaughan, 1988). Both the A and B subunits of CT and LT are synthesized intracellularly as precursor proteins which, after removal of the leader peptide and translocation across the cytoplasmic membrane, assemble in the periplasmic space to form the final AB5 complex. While V. cholerae exports the CT toxin into the culture medium, LT remains

Fig. 33. X-ray structure of heat-labile enterotoxin LT of E. coli. The catalytic domain A1 is yellow, the linker domain A2 is blue, and the five monomers of the B subunit are all represented in different colors.

associated to the outer membrane bound to lipopolysaccharide (LPS; Horstman et al., 2002). The corresponding genes of CT and LT are organized in a bicistronic operon and are located on a filamentous bacteriophage and on a plasmid, respectively (So et al., 1978). The A subunit (Fig. 34, left panel) is a 27-kDa monomer composed of a globular structure and linked to the B domain by a trypsin-sensitive loop and a long a-helix, which inserts inside the core of the B pentamer thus anchoring the two subunits. For full activity, the A subunit needs to be proteolytically cleaved and reduced at the disulfide bridge between cysteines 187 and 199 to give two fragments: the enzymatic subunit A1 and the linker fragment A2 (Lai et al., 1981).

CHAPTER 1.28

Bacterial Toxins

921

Fig. 34. Left side: front view of the catalytic A subunit, with the toxic moiety A1 in pale green and the linker domain A2 in violet; cysteines 187 and 199 involved in the disulfide bridge are red. Right side: bottom view of the pentameric receptor-binding domain B.

Whereas in cholera toxin the proteolytic process is performed during biosynthesis by an endoprotease (Booth et al., 1984), in the case of LT, it occurs by extracellular processes; in both cases, the reduction is thought to take place at the surface of the target cell. The enzymatically active domain A binds NAD and transfers the ADP-ribose group to an Arg residue located within the central portion of several GTP-binding proteins such as Gs, Gt and Golf. Upon ADP-ribosylation of Gs, in particular, the adenylate cyclase is permanently activated, causing an abnormal intracellular cAMP accumulation, which in turn alters ion transport and thus is the main reason for the toxic effects (Field et al., 1989a, 1989b). A peculiar feature of CT and LT is that the basal ADP-ribosyltransferase activity is enhanced by interaction with 20-kDa guaninenucleotide binding proteins, known as “ADPribosylation factors” (ARFs; Tsai et al., 1988; Moss and Vaughan, 1991). After receptor binding, the holotoxins are internalized and undergo retrograde transport through the Golgi to the endoplasmic reticulum (ER). Recent studies show that both A and B subunits move together from the cell surface into the ER, and this depends on the B-subunit binding to ganglioside GM1. The KDEL motif in the A2 chain does not appear to affect retrograde transport, but slows recycling of the B-subunit from ER to distal Golgi stacks. Specificity for GM1 in this trafficking pathway is shown by the failure of the E. coli type II toxin LTIIb that binds ganglioside GD1a to concentrate in lipid rafts, enter the ER, or induce toxicity. These results show that the B subunit carries the A1 chain from cell surface into the ER where they dissociate, and that a membrane lipid with strong affinity for lipid rafts provides the dominant sorting motif for this pathway (Fujinaga et al., 2003). In the ER, the A1-chain of the CT unfolds and enters the cytosol by a process termed “retro-translocation.”

Upon entering the cytosol, the A1-chain rapidly refolds, binds ARF and induces toxicity (Lencer et al., 1995). The B subunits persist in the Golgi and are subsequently degraded. The exact localization of the ARF-binding site is still unknown, but it has emerged from recent studies that the two domains (the NAD-binding and ARF-binding) are independent and located in different regions of the A domain (Stevens et al., 1999). When the toxins are released in the intestine during infection, the major consequence is intestinal fluid accumulation and watery diarrhea (also typical symptoms of the diseases; Holmgren, 1981). The B domain (Fig. 34, right) is composed of five identical subunits (each 11.5 kDa) that are arranged in a symmetric shape around a central pore inside which the C-terminal portion of the catalytic domain (A2) is inserted (Sixma et al., 1991, 1993). Their secondary structure consists predominantly of two three-stranded antiparallel b-sheets, a short N-terminal helix, and a long central helix. Although still well conserved in terms of quaternary structure, CT and LT B domains have a lower degree of primary sequence homology than the corresponding A domains. Interestingly, the B subunit of LT-II, although maintaining a conserved structure, lacks any sequence homology with the corresponding B domains of CT and LT-I (Domenighini et al., 1995). In addition to their function as receptorbinding domains and as carriers of the toxic moieties, the B subunits possess specific biological activities such as induction of apoptosis of CD8+ and CD4+ T cells (Truitt et al., 1998) and the property to function as potent mucosal adjuvants (Xu-Amano et al., 1994). This feature has been extensively used to develop mucosal vaccines against cholera and ETEC infection, and to induce a mucosal response also against the other antigens used.

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wild type

LT K63

To produce molecules suitable for these pharmacological applications but completely devoid of toxic activity, more than fifty mutant derivatives have been constructed both for the A and B subunits. Among those which completely inactivate the toxin, the best characterized are LTK63 (Fig. 35), LT-K97 and LT-K7, all in the vicinity of the catalytic domain, and for which the 3D structures have also been determined (Merritt et al., 1995; Van den Akker et al., 1995, 1997). In the case of LT-K63 (and the corresponding CTK63), where the wildtype Ser in position 63 is substituted with a Lys, the mutated proteins are enzymatically inactive and nontoxic, either in vitro and in vivo, but are otherwise indistinguishable from the wildtype. In fact, they are still able to bind the receptor and the ARFs (Stevens et al., 1987), and the crystal structure and that of wildtype LT are almost perfectly superimposable except for the catalytic site, where the bulky side-chain of Lys-63 fills the catalytic pocket thus making it unsuitable for NAD entrance and binding (Giannelli et al., 1997; Douce et al., 1998). Another interesting mutant is LT-K97, where the substitution Val/Lys introduces a salt bridge between Lys-97 and the carboxylate of Glu-112, thus making it unavailable to further interactions. This observation suggests a dominant role of this glutamic acid in the enzymatic reaction. Mutations affecting the B domain lead often to products that can no longer bind to eukaryotic receptors, as is the case of LTB-D33, which contains a glycine-to-aspartic acid substitution in position 33. These types of mutants have been found to be almost completely nonimmunogenic at mucosal surfaces, suggesting that an intact receptor-binding site is necessary not only for binding but also for immunogenicity and adjuvanticity (Guidry et al., 1997).

Clostridium perfringens Alpha-Toxin This toxin (Fig. 1, panel 16) is the most important toxin produced by Clostridium perfringens and is responsible for gas gangrene or clostridial myo-

CHAPTER 1.28 Fig. 35. Three-dimensional structure of the enzymatic cavity of the wildtype LT (left) and of the mutant LT-K63 (right). The arrows point out how much a single amino acid substitution can affect the dimension of the pocket and thus the entrance of NAD.

necrosis (Stevens et al., 1987; Florez-Diaz et al., 2003). It plays a key role in the spread of the infection either by suppressing host immune responses, triggering the release of inflammatory mediators, or causing changes in intracellular calcium levels. Specific mutants of C. perfringens that do not produce the toxin are unable to cause disease, and vaccination with a genetically engineered toxoid has been shown to induce protection against gas gangrene (Williamson and Titball, 1993). This virulence factor is a 370-amino acid zinc metalloenzyme that also displays phospholipase C (PLC) activity (Leslie et al., 1989); nevertheless, not all the bacterial PLCs act as virulence determinants, therefore this enzymatic activity is not sufficient for toxicity. Alpha-toxin is capable of binding to mammalian cell membrane and cleaving membranebound phosphatidylcholine (or sphingomyelin) to produce phosphocholine and diacylglycerol (or ceramide). The reaction product diacylglycerol, which is a leukotriene precursor, is believed to be the responsible of the subsequent lethal effects. The crystal structure of a-toxin has been recently solved (Naylor et al., 1998; Fig. 36), indicating the presence of two distinct domains in the molecule. Whereas the N-terminus is mainly organized as a globular a-helical domain that contains the active site, the b-sandwich C-terminal subunit is involved in membrane binding and shows strong structural analogy to eukaryotic calcium-binding C2 domains. A flexible linker containing a series of highly mobile residues connects the two domains. In addition, the C-terminal subunit displays hemolytic and sphingomyelinase activities and primarily contributes to the toxin’s lethal effect, even if it is completely devoid of toxic activity when used alone. Nevertheless, immunization with this domain affords full protection from disease in mouse models, thus indicating that the protective epitopes are located in this portion of the molecule (Titball et al., 1993; Nagahama et al., 2002).

CHAPTER 1.28

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acterized by a highly repetitive domain called the “clostridial repetitive oligopeptide” (CROP), identified as the site of interaction with a carbohydrate structure as well as the ligand to which neutralizing antibodies bind (von EichelStreiber, 1994). A central hydrophobic region contains several predicted transmembrane segments and is believed to function as the translocation unit.

Bordetella pertussis Adenylate Cyclase

Fig. 36. Three-dimensional structure of Clostridium perfringens a-toxin. The N-terminal and C-terminal domains are green and violet, respectively, and the flexible linker is orange.

Recently, other bacterial PLCs, like those from L. monocytogenes and Mycobacterium tuberculosis, have been implicated in the pathogenesis of a number of diseases (Wadsworth et al., 1999; Raynaud et al., 2002).

Clostridium difficile Toxins A and B Enterotoxin A (TcdA) and cytotoxin B (TcdB) of Clostridium difficile are the two virulence factors responsible for the induction of antibioticassociated diarrhea. These toxins have molecular masses of 308 and 270 kDa, respectively, and belong to the class of large clostridial cytotoxins (Lyerly et al., 1986; Knoop et al., 1993). The toxin genes tcdA and tcdB together with three accessory genes (tcdC–E) constitute the pathogenicity locus (PaLoc) of C. difficile (Cohen et al., 2000). Primary sequence homology between tcdA and tcdB gene products is higher than 60% identity (von Eichel-Streiber et al., 1994). Upon binding to eukaryotic cells and translocation across membranes via receptor-mediated endocytosis, TcdA and TcdB monoglucosylate small GTP-binding proteins such as Rho, Rac and Cdc42 at a threonine residue (Just et al., 1995a, 1995b; Ciesla and Bobak, 1998). In most cells, C. difficile toxins induce depolymerization of the actin cytoskeleton, leading to a morphology similar to that induced by C3-like transferases. While toxin B has potent cytotoxic activity in vitro, the enterotoxic activity of C. difficile in animals has been mainly attributed to toxin A. From the structural point of view, they are composed of two portions: the N-terminal nonrepetitive two thirds corresponding to the catalytic subunit, and the C-terminal third char-

Adenylate cyclase (CyaA) is a toxin produced by Bordetella pertussis, B. bronchiseptica and B. parapertussis (Weiss and Hewlett, 1986). It is essential in the early stages of bacterial colonization of the respiratory tract and can induce apoptosis of lung alveolar macrophages (Goodwin and Weiss, 1990; Khelef et al., 1993). Organized as a bifunctional protein, CyaA (177 kDa) is composed of an N-terminal cellinvasive and calmodulin-dependent adenylate cyclase domain (residues 1–400) fused to a poreforming hemolysin (residues 401–1706; Glaser et al., 1988; Bejerano et al., 1999; see also the section Pore-Forming Toxins: RTX Hemolysins). Unlike most of the other members of the RTX family that are secreted into the supernatant, CyaA remains associated to the bacterial surface, through interactions with filamentous hemagglutinin (FHA). This toxin forms small cation-selective channels in lipid bilayer membranes and delivers into the cytosol of target cells the adenylate cyclase (AC) domain, which, upon binding to calmodulin, catalyzes an uncontrolled conversion of ATP to cAMP, thus causing intoxication and disruption of cellular functions (Ladant and Ullmann, 1999). Calcium has been shown to play a fundamental role in channel formation (Knapp et al., 2003). Furthermore, it was also demonstrated that the ability of the AC domain to form pores and translocate across the membrane is strictly linked to the correct folding of an amphipathic a-helix spanning residues 509–516. Substitution of Glu-509 with a helixbreaker proline residue, in fact, significantly reduced the capacity of the toxin to undergo translocation (Osickova et al., 1999). A very similar function and mechanism of action is that of ExoY, an adenylate cyclase produced by Pseudomonas aeruginosa and injected into the cytoplasm of eukaryotic cells by the type III secretion apparatus (see Table 1, and the section Toxins Injected into Eukaryotic Cells in this Chapter). However, differently from CyaA, ExoY is not activated by calmodulin. In vivo, following infection with ExoY-expressing strains, CHO cells showed a rounded morphology, which correlated with increased cAMP levels (Yahr et al., 1998).

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CHAPTER 1.28

Anthrax Edema and Lethal Factors Lethal factor (LF) and edema factor (EF) proteins, produced by Bacillus anthracis, combine with the protective antigen PA to give the lethal (PA+LF) and edema (PA+EF) toxins (Brossier et al., 2000; Collier and Young, 2003; Fig. 1, panel 17). In both complexes, the PA has the pore-forming, receptor-binding activity (see the section PoreForming Toxins in this Chapter), whereas EF and LF display, in turn, the toxic activities. The EF and LF genes are located on a large plasmid (Mikesell et al., 1983) and encode precursors of approximately 800 residues. Cleavage of the N-terminal signal peptides yields mature EF and LF proteins with molecular masses of 88.8 kDa and 90.2 kDa, respectively. These virulence factors enter cells by binding to proteolytically activated, receptor-bound, oligomeric PA; following receptor-mediated endocytosis, the low pH causes a conformational change in PA, allowing the translocation of EF-LF across cell membrane (Collier, 1999). The EF-LF is then endocytosed and translocated from endosomes directly to the cytosol of cells, where both toxins perform their toxic activities (Fig. 37). The binding sites of EF and LF on PA have been recently mapped (Cunningham et al., 2002). Once inside the cells, EF binds calmodulin and catalyzes an unregulated production of the second messenger cAMP, thereby perturbing the normal cell regulatory mechanisms (Goyard et al., 1989). Calcium influx is required for inducing cyclic AMP toxicity in target cells (Kumar et al., 2002). Whereas the PA-binding domain displays a strong sequence homology to lethal factor LF, the catalytic domain is more similar to the other known adenylate cyclase CyaA toxin of Bordetella pertussis (Escuyer et al., 1988). On the other hand, LF cleaves the amino-terminus of the cellular mitogen-activated protein kinase kinases (MAPKK1 and MAPKK2), thus causing inhibition of the MAPK signal transduction pathway, which is key to cellular proliferation and signal transduction processes in the cell (Duesbery et al., 1998; Vitale et al., 1999). Recently, the 3D structures of LF and EF have been solved (Fig. 38). LF comprises four domains: domain I binds the membranetranslocating component of anthrax toxin (PA); domain II resembles the ADP-ribosylating toxin from Bacillus cereus; domain III is inserted into domain II, and seems to have arisen from a repeated duplication of a structural element of domain II. Domain IV is distantly related to the zinc metalloprotease family, and contains the catalytic center (Pannifer et al., 2001). The catalytic portion of EF is made by three globular domains. The active site is located at the inter-

PA PA binds to receptor Receptor Protease

20

Protease cleaves

63

20-kDa fragrnent released EF and LF sites exposed

Receptor Protease

EF

LF

EF/LF 63

EF or LF binds

Receptor

EF or LF enters endosome

EF/LF 63

Receptor

EF or LF translocates from endosome to cytosol H+

Then, EF binds Colmodulin and makes cAMP. while LF cleaves MAPKK1 and MAPKK2 Cdr EF LF ATP

cAMP

MAPKK1 and MAPKK2

Fig. 37. Mechanism of PA-mediated entry and intoxication of anthrax LF and EF toxins.

face of two domains (CA and CB), which together form the catalytic core, containing the catalytic residue His351. EF has been crystallized both alone and in complex with calmodulin. The differences between the two forms are induced by calmodulin, which acts by stabilizing the conformation of the substrate-binding-site of EF (Drum et al., 2002). Interestingly, a remarkable level of primary sequence similarity can be detected between EF and the N-terminal, calmodulin-binding domain of Bordetella adenylate cyclase CyaA. In particular, His351 is conserved between the two proteins. Once in the cytoplasm, LF acts as a zincmetalloprotease disrupting normal homoeostatic functions. The macrophage is a uniquely sensitive cell type that seems to be a vital global mediator of toxin-induced pathologies. Removal of macrophages from mice renders them insensitive to LF challenge (Hanna, 1999). In addition, LF, but not EF, is able to cause apoptosis in human endothelial cells. As a con-

CHAPTER 1.28

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925

Domain I

Helical domain

Domain II

CB

Domain IV

CA

Domain III

PANEL A

PANEL B

Fig. 38. Crystal structures of the catalytic portion of anthrax lethal factor (panel A) and edema factor in complex with calmodulin (panel B). Panel A. The four domains are in different colors. The zinc atom complexed by domain IV is indicated by an arrow.

sequence, the observed endothelial toxicity contributes to vascular pathology and hemorrhage during systemic anthrax (Kirby, 2004).

E. coli Cytotoxin Necrotizing Factors Cytotoxin necrotizing factors (CNF1 [Fig. 1, panel 18] and CNF2), single-chain proteins of 115 kDa produced by a number of uropathogenic and neonatal meningitis-causing pathogenic E. coli strains (Caprioli et al., 1984; De Rycke et al., 1987), are immunologically related and share 85% identity. They also share some similarity with the dermonecrotic toxin of Pasteurella multocida and Bordetella pertussis (Schmidt et al., 1999). Both CNF1 and CNF2 toxins are encoded by a single structural gene with a low G+C content (35%). However, whereas cnf1 is chromosomally encoded, cnf2 is carried on a large transmissible F-like plasmid called “Vir” (Oswald and De Rycke, 1990; Falbo et al., 1992). These toxins induce ruffling, stress fiber formation, and cell spreading in cultured cells by activating the small GTP-binding proteins Rho, Rac and Cdc42, which control assembly of actin stress fibers (Oswald et al., 1994). CNF1 induces only a transient activation of Rho GTPase and a depletion of Rac by inducing the addition of an ubiquitin chain, which is known to drive to specific degradation by the proteasome. Reduction of Rac GTPase levels induces cell motility and

Fig. 39. Crystal structure of the active site of E. coli CNF1. The catalytic site composed by Cys866-His881 is colored in blue.

cellular junction dynamics allowing efficient cell invasion by uropathogenic bacteria (Doye et al., 2002). The catalytic region of CNF1 has been crystallized (Buetow et al., 2001; Fig. 39). The active site contains a catalytic triad, which is positioned in a deep pocket, thus explaining the restricted access to unspecific substrates and therefore its specificity. Very likely, some type of conformational rearrangement is required also to accomodate Rho in this narrow cavity.

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Recently, a CNF1-like toxin (CNFY) has been identified also in Yersinia pseudotubercolosis (Lockman et al., 2002). Differently from the E. coli CNFs, CNFY has been shown to selectively activate RhoA (Hoffman et al., 2004).

Bordetella Dermonecrotic Toxin Dermonecrotic toxin (DNT) is produced by Bordetella species as a single-chain polypeptide chain of 1464 amino acids, which is composed of a C-terminal portion that contains the catalytic site, and of an N-terminal receptor-binding domain. DNT shares about 30% identical residues in the catalytic domain with E. coli CNF1, including the catalytic Cys and His residues. DNT is a transglutaminase, which catalyzes the deamidation or polyamination at Gln63 of Rho and of the corresponding residues of Rac and Cdc42 (Horiguchi, 2001). This activity causes alteration of cell morphology, reorganization of stress fibers, and focal adhesions on a variety of animal models. Recently, it has been demonstrated that the initial 54 amino acids of DNT are sufficient for cell surface recognition. However, the receptor is still unknown.

Cytolethal Distending Toxins The cytolethal distending toxin (CDT) produced by Haemophilus ducreyi (HdCDT) is the prototype of a growing family of bacterial toxins that act by inducing cell enlargement followed by cell death (Cortes-Bratti et al., 1999; Frisan et al., 2003). HdCDT is a complex of three proteins (CdtA, CdtB and CdtC) encoded by three genes that are part of an operon. Members of this family have been identified in E. coli, Shigella, Salmonella, Campylobacter, Actinobacillus and Helicobacter hepaticus (Okuda et al., 1995; LaraTejero and Galan, 2001; Hoghjoo et al., 2004; Pickett et al., 2004; Shenker et al., 2004; Young et al., 2004). The overall sequence similarity varies among the different members of this family of toxins. HdCDT intoxicates eukaryotic cells by causing a three- to fivefold gradual distension and induces cell cycle arrest in the G2 phase. It has also been shown to induce DNA doublestrand breaks and formation of actin stress fibers via activation of the small GTPase RhoA. Recently it has been shown that CdtB is the active subunit of the CDT toxin and acts as a nuclease. All the amino acids predicted to be important for nuclease activity are conserved in the CdtB of different bacteria, suggesting that the mechanism of action is the same for all CDT toxins. On the other hand, CdtA and CdtC are able to bind to the surface of HeLa cells, therefore playing a role in the delivery of the active domain to target cells (Lee et al., 2003).

CHAPTER 1.28

Toxins Acting on the Cytoskeleton Structure The cytoskeleton is a cellular structure that consists of a fiber network composed of microfilaments, microtubules, and the intermediate filaments. It controls a number of essential functions in the eukaryotic cell and participates in all kinds of cellular movement and transport; furthermore, the cytoskeleton is involved in processes like exo- and endocytosis, vesicle transport, cell-cell contact, and mitosis (Kabsch and Vandekerckhove, 1992). The group of cytoskeleton-affecting bacterial toxins comprises not only virulence factors that directly act on particular elements of the cytoskeleton, but also proteins that perform an indirect action by affecting regulatory components, which control its organization (Aktories, 1994; Richard et al., 1999). Most of them do it by modifying the regulatory, small G proteins, such as Ras, Rho, Rac and Cdc42, which control cell shape. These toxins, which have a dramatic but indirect effect on the cytoskeleton and are described in the section Toxins Acting on Signal Transduction, are E. coli CNF and C. difficile enterotoxins A and B. Other toxins acting on regulatory G proteins are exoenzyme S, C3 and YopE, which are described below as toxins that are directly injected into the eukaryotic cells. Other bacterial molecules that cannot be strictly considered toxins but that have a powerful ability to polymerize actin are ActA and IcsA of Listeria and Shigella, respectively. These are described elsewhere in this volume (see Listeria and Relatives in Volumn 4 and The Genus Shigella in Volumn 6). Another toxin acting indirectly on the cytoskeleton is the zonula occludens toxin (Zot) produced by V. cholerae, a toxin with an unknown mechanism of action that modifies the permeability of tight junctions (Zot is described in the paragraph Toxins with Unknown Mechanism of Action in this Chapter). In the following section we consider only toxins that have the cytoskeleton as a direct target. The only toxin shown to affect directly the cytoskeleton is the C2 toxin of C. botulinum, which ADPribosylates monomeric actin, making it unable to polymerize. A second protein that has recently been described as being able to bind actin and stabilize the fibers supporting the ruffles induced by the Salmonella type III secretion system is SipA (described in the section Toxins Injected into Eukaryotic Cells in this Chapter). Representatives of both subgroups can be identified among the class of ADP-ribosylating factors that ultimately display their toxic effect on the cytoskeleton of eukaryotic cells. In fact, whereas the family of Clostridium botulinum toxin C2, clostridial toxin C3 (and related pro-

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teins), and Pseudomonas aeruginosa exoenzyme S (Exo S) act on small GTP-binding proteins that regulate the correct functioning of the cytoskeleton, and thus have an indirect toxic effect (Coburn et al., 1999).

Clostridium botulinum Toxin C2 and Related Proteins Clostridium botulinum toxin C2 is the main representative of a class of binary cytotoxins produced by clostridial species that predominantly act on polymerized actin microfilaments of 7– 9 nm in diameter (Aktories et al., 1986; Aktories and Wegner, 1992). C2 ADP-ribosylate monomeric G-actin at an arginine residue (Aktories et al., 1986; see the section ADP-Ribosylating Toxins in this Chapter). Because this arginine (Arg-177) is a contact site between actin monomers, the binding of the ADP-ribose moiety prevents actin’s polymerization. Other members of this family are C. perfringens iota toxin (Stiles and Wilkins, 1986; Perelle et al., 1993) and the related C. spiroforme and C. difficile ADP-ribosylating toxins (Popoff and Boquet, 1988a; Just et al., 1994), which are generally classified as iota-like toxins. These binary toxins are constructed according to the A/B model architecture, but in this case the two domains reside in separate molecules that interact to cause the toxic effect. Therefore, these toxins have an enzymatically active and toxic domain (A) and a binding component (B), which is essential for the binding at the cell surface and for the translocation inside the cell. Clostridium botulinum toxin C2 is an extremely toxic agent, which induces hypotension, increase in intestinal secretion, vascular permeability, and hemorrhaging in the lungs. In contrast to botulinum neurotoxins, C2 does not seem to display any neurotoxic effect. The two molecules that constitute its toxic moiety are classified as C2-II (for the binding component) and C2-I (for the enzymatic component). The C2-II is a 100-kDa protein that must be proteolytically cleaved to a 75-kDa fragment before it can bind to the surface receptor; upon this interaction, a binding site for the 50-kDa C2-I component is activated and the toxic domain is taken up by receptor-mediated endocytosis (Ohishi, 1987). Substrates of the C2-I toxin are b/g-non-muscle actin and g-smooth muscle actin, but not a-actin isoforms. Conversely, the related iota toxin of Clostridium perfringens has been found to ADP-ribosylate all actin isoforms (Mauss et al., 1990). The iota toxin is a binary toxin produced by Clostridium perfringens type E, which has been implicated in fatal calf, lamb and guinea pig enterotoxemias (Madden et al., 1970). Structurally, it has two independent

Fig. 40. Crystal structure of the catalytic domain C2I of C. perfringens C2 toxin (red and yellow) in complex with NADH (pink).

domains: Ia, which is the ADP-ribosyltransferase, and Ib, which is involved in the binding and internalization of the toxin by the cell (Stiles and Wilkins, 1986). The crystallization of the C2I component in complex with its substrate NADH has recently been achieved (Tsuge et al., 2003; Fig. 40), showing a close relationship of iota toxin with insecticidal protein VIP2 of Bacillus cereus. Clostridium difficile induces its pathogenic effects by secreting a number of potent cytotoxins; one, in particular, has been found to possess ADP-ribosyltransferase activity (CDT). CDT acts on the cytoskeleton structure by disaggregating actin filaments and thus provokes an increase of globular actin (G-actin; Popoff et al., 1988b; Gulke et al., 2001). Another member of the group of iota-like toxins is the Clostridium spiroforme toxin, composed of a toxic subunit Sa and a binding subunit Sb (Popoff et al., 1989). The level of primary sequence homology detected among the enzymatic and binding components of this class of ADP-ribosylating toxin ranges from 32% to 80% identity, the binding domains being the better conserved. The C2 toxin is the one with the lower degree of sequence conservation, and this correlates with the fact that it does not appear to be crossreactive with the other iota-like toxins. Experiments of site-directed mutagenesis have helped to define for these toxins an active site very similar to those described for the better studied members of the family of ADP-ribosyltransferases (Barth et al., 1998; see the section ADP-Ribosyltransferases: A Common Structure of the Catalytic Site in this Chapter).

Escherichia coli Lymphostatin Lymphostatin is a very recently identified protein in enteropathogenic strains of E. coli (EPEC; Klapproth et al., 2000).

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A leading cause of diarrhea among infants in developing countries, EPEC is also one of the few known bacterial causes of chronic diarrhea. These strains are characterized by their ability in host cells to induce cytoskeletal rearrangements that result in the formation of adhesion pedestals. This mechanism known as “the attaching and effacing effect” (Moon et al., 1983; Khoshoo et al., 1988) ultimately allows the bacterium to colonize the host for prolonged periods. Lymphostatin also has been identified as one of the primary factors that selectively block the production of interleukin-2 (IL-2), IL-4, IL-5 and g interferon by human peripheral cells and inhibit proliferation of these cells, thus interfering with the cellular immune response (Klapproth et al., 1995). Lymphostatin, a very large toxin with a predicted molecular weight of 366 kDa, shares significant homology with the catalytic domain of the large clostridial cytotoxins, including toxins A and B of Clostridium difficile, lethal toxin of C. sordelii, and a toxin of C. novyi. Its corresponding gene, lifA, with 9669 bp, is the largest reported gene in E. coli. Some lifA mutants of EPEC have been constructed to verify the lymphocyte inhibitory factor (LIF) activity of its gene product; lysates of this mutant lacked the ability of wildtype EPEC lysates to inhibit expression of IL-2, IL-4 and g interferon mRNA and protein in mitogen-stimulated lymphocytes, while the expression of IL-8 was unaffected (Klapproth et al., 2000). Experiments of colony hybridization performed using an internal fragment of the lifA gene identified a similar gene present in most of the EPEC and enterohemorrhagic E. coli (EHEC) strains able to produce the attaching and effacing lesions on host epithelial cells, but this gene was not found in other

E. coli and related organisms (Klapproth et al., 2000).

Toxins Acting on Intracellular Trafficking Vesicle structures are essential in the eukaryotic cell for a number of vital processes such as receptor-mediated endocytosis and exocytosis; these are used either to internalize portions of the plasma membrane and address them to the specialized compartment, or to transport to the cell surface molecules synthesized in the ER and modified in the Golgi apparatus. One example of exocytic pathway is that involving the release of neurotransmitters that are contained within small synaptic vesicles packed at synaptic terminals; the majority of these vesicles are bound to the cytoskeleton and are not directly available for immediate release, but some of them are present at the cytosolic face of the presynaptic membrane and are ready to release their content. However, at low calcium concentrations, only an occasional vesicle fuses to the presynaptic membrane, giving rise to a depolarization event. This event leads to the opening of calcium channels and thus to an increase of calcium concentration, which finally triggers the fusion of the neurotransmitter vesicles with the plasma membrane. Recently, this field was greatly advanced by the identification of the eukaryotic molecules responsible for vesicle docking and membrane fusion. Three of these proteins (namely vesicleassociated membrane protein [VAMP]/ synaptobrevin, synaptosome-associated protein [SNAP-25], and syntaxin) are the specific targets of a number of neurotoxins produced by bacteria of the genus Clostridium (CNTs; Montecucco and Schiavo, 1994; Fig. 41).

Presynaptic cell Zn

NT

Cleaves

NT

Synaplobrevin (v-SNARE) NT NT

Zn

Syntaxin SNAP-25

Cleaves

NT

l-SNARE –Toxin

NT release

+Toxin

Fig. 41. Mechanism of action of clostridial neurotoxins.

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The CNT family is composed of tetanus neurotoxin (TeNT) and seven serotypes of botulinum neurotoxins (BoNT/A–BoNT/G), which are specific zinc-dependent proteases whose action finally causes the block of neuroexocytosis (Schiavo et al., 1992; Pellizzari et al., 1999; Lalli et al., 2003). The degree of sequence homology detected among this group of toxins is high, ranging from 30% to more than 50% identity.

Clostridium tetanii Neurotoxin Tetanus neurotoxin (TeNT; Fig. 1, panel 19) is the unique causal agent of the pathological condition of spastic paralysis known as tetanus. This is one of the most potent toxins known so far, with a 50% lethal dose (LD50) in humans of 0.1– 1.0 ng/kg. The TeNT is produced by Clostridium tetanii as a single chain polypeptide of 150 kDa that, following proteolytic cleavage, is divided into fragments H (heavy) and L (light) held together by a disulfide bridge. Its overall structure is similar to that of A/B toxins, where the toxic subunit A is represented here by the light chain L, and subunit B is constituted by the HC and HN domains. The heavy chain is composed of fragments HC, which has recently been found to bind di- and trisialylgangliosides on neuronal cell membranes, and HN, which is involved in the transmembrane translocation of the L chain to the cytosol (Schiavo et al., 1990; Shapiro et al., 1997). The L chain is a 50-kDa fragment containing the –HExxH– motif typical of metalloproteases. It binds zinc and specifically cleaves VAMP/synaptobrevin, a eukaryotic factor essential for membrane fusion (Rossetto et al., 1995). The first step of intoxication is the specific binding of domain HC of TeNT to both high and low affinity receptors exposed on the presynaptic neuronal membrane at neuromuscular junctions (Montecucco, 1986); the second step is internalization of TeNT into the peripheral motoneuron and then retrograde axonal transport. The TeNT is released through the postsynaptic membrane into the synaptic space where it enters into the inhibitory interneurons of the central nervous system through receptor-mediated endocytosis (Halpern and Neale, 1995). At this point, while the HC domain is in the vesicle, the translocation domain HN helps the catalytic light chain L to cross the vesicle membrane and gain access to the cytosolic compartment where L performs its toxic activity on VAMP/synaptobrevin (Montal et al., 1992). Interestingly, domain HC retains the unique transport properties of the intact holotoxin and is capable of eliciting a protective immunological response against the full-length tetanus neurotoxin. A single zinc atom is bound to the L chain

Fig. 42. Crystal structure of the receptor-binding domain HC of tetanus neurotoxin in complex with a ganglioside analogue (in red). The N-terminus and C-terminus are colored in blue and green, respectively. The residues probably involved in ganglioside binding are yellow (see Fig. 1, panel 18).

of TeNT and is essential for toxicity. This specific metallo-dependent proteolytic activity is common to the other clostridial toxins and to the lethal factor (LF) of Bacillus anthracis. The crystal structure of the receptor-binding fragment HC of tetanus neurotoxin has been recently determined at 2.7 Å resolution (Umland et al., 1997; Fig. 42) revealing an N-terminal jellyroll domain and a C-terminal b-trefoil domain. To determine which amino acids in tetanus toxin are involved in ganglioside binding, homology modeling was performed using recently resolved X-ray crystallographic structures of the tetanus toxin HC fragment. On the basis of these analyses, the amino acids tryptophan 1288, histidine 1270, and aspartate 1221 were found to be critical for binding of the HC fragment to ganglioside GT1b (Fotinou et al., 2001; Louch et al., 2002). Although the overall sequence homology detected among clostridial neurotoxins is significant, this similarity weakens in the region encompassing the C-terminal domain (Murzin et al., 1992); the fact that each toxin possesses its own unique receptor and is immunologically distinct from the others has been attributed to sequence divergence of this domain which, therefore, could be responsible for receptor specificities (Lacy et al., 1999).

Clostridium botulinum Neurotoxins These neurotoxins (BoNT/A-G; Fig. 1, panel 20) are the causative agents of the flaccid paralysis typical of clinical botulism intoxication (Hatheway, 1995). All of them are zinc-dependent proteases that show a strong tropism for the neuromuscular junction (Simpson, 1980; Rossetto et al., 1995), where they bind to still unidentified receptors in a strictly serotype-specific manner. This binding step is followed by the entry of the toxin into the cytoplasm of the motoneurons and

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by specific proteolytic cleavage of intracellular targets belonging to the family of soluble Nethylmaleimide-sensitive, fusion factor attachment protein receptors (SNARE). Four out of the seven botulinum neurotoxins (BoNT/B, D, F and G) cleave VAMP/synaptobrevin, another two act specifically on SNAP/25, whereas the last one, BoNT/C, cleaves both syntaxin and SNAP/ 25 substrates. In all cases, the ultimate effect is the total block of acetylcholine release (Montecucco and Schiavo, 1995). These toxins are generally produced as large complexes of 300–900 kDa containing additional proteins such as hemagglutinin (300 kDa) and nontoxic peptides, which are believed to act as stabilizing agents of the neurotoxins in the gut environment (Sakaguchi, 1983). The BoNTs are synthesized as inactive polypeptide chains of 150 kDa, which (following proteolytic cleavage) divide into two chains of 50 and 100 kDa that remain linked by a disulfide bridge. The catalytic function is carried by the 50kDa fragment, the light chain L (residues 1–437), whereas the 100-kDa subunit (heavy chain, H) contains both the translocation (residues 448– 872) and the receptor-binding domains (residues 873–1295; Krieglstein et al., 1994). The crystal structure determined for the full-length polypeptide of BoNT serotype A (Lacy et al., 1998; Fig. 43) reveals a number of remarkable features, particularly related to the peculiar structure of the translocation domain. This contains, in fact, a central pair of a-helices 105 Å long and a 50-residue loop that wraps around the catalytic domain in a belt-like fashion, partially occluding the activesite pocket. This unusual loop bears the site of the proteolytic cleavage, which is required for

Fig. 43. X-ray structure of Clostridium botulinum neurotoxin serotype A. The 50-kDa catalytic domain (L) is colored in yellow, with the zinc-binding domain in green. The Nterminal portion of the 100-kDa subunit involved in translocation is blue, whereas the C-terminal receptor-binding moiety is in magenta. The disulfide bond linking the two 50and 100-kDa fragments is colored in red (see Fig. 1, panel 20).

CHAPTER 1.28

activation of the toxin; the fact that in the protoxin, the translocation domain shields the active site explains why the catalytic activity in test tube experiments is greatly enhanced by reduction of the disulfide bond. The fold of the translocation domain suggests a mechanism of pore formation different from that displayed by other poreforming toxins. The helices are antiparallel and amphipathic and twist around each other in a coiled-coil-like structure. In addition, the domain has two strand-like segments that lie parallel to the helical axis and are predicted to be directly involved in membrane spanning. Very recently, the X-ray structure obtained for the recombinant form of chain L of BoNT-A has shed light on a possible novel mode of substrate binding and catalytic mechanism (Segelke et al., 2004). The highest degree of homology detected among this family of clostridial neurotoxins is concentrated in the light chain L (30–60% identity; particularly its N-terminus), probably involved in substrate recognition, and in the central portion that contains the catalytic zincbinding motif –HexxH– characteristic of zinc endopeptidases. The zinc atom coordinated by this pocket is required for the in vivo toxicity of BoNTs. Years ago, medical experiments demonstrated that injection of BoNT/A is very effective in strabismus; since then, the therapeutic applications of these neurotoxins have been extended to a variety of diseases which benefit from a functional paralysis of the neuromuscular junction, and all the BoNTs are under clinical testing (Jankovich and Hallett, 1994).

Helicobacter pylori Vacuolating Cytotoxin Vac A Highly pathogenic strains of Helicobacter pylori, the etiological agent of peptic ulcer and gastritis (Cover and Blaser, 1992), produce vacuolating cytotoxin A (VacA; Papini et al., 1994; Fig. 1, panel 21). This toxin is responsible for massive growth of vacuoles within epithelial cells and, when administered to mice, VacA causes loss of gastric gland architecture, cell necrosis, and gastric ulceration (Telford et al., 1994). Synthesized as a 140-kDa precursor, VacA is secreted from the bacterium through its 45-kDa carboxy-terminal domain, using a mechanism similar to that of neisserial IgA proteases (Schmitt and Haas, 1994; Fiocca et al., 1999). When purified from the culture supernatant of Type I H. pylori strains, the protein has a molecular weight of approximately 600–700 kDa, suggesting the idea of a multimeric complex; electron microscopy studies have in fact demonstrated the flower-shaped structure of the toxin (Lupetti et al., 1996; Fig. 44) resulting from the aggregation of either six or seven monomers, each

CHAPTER 1.28

Bacterial Toxins

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Fig. 44. Vacuolating cytotoxin structure: heptameric and hexameric forms of VacA as observed in electron micrographs of quick-freeze, deepetched preparations. The oligomers are approximately 30 nm in diameter with a 10–12 nm central cavity.

comprising the 95-kDa amino-terminal region of the VacA precursor. Recently, a model has been proposed to show how VacA can insert into membranes forming hexameric, anion-selective pores (Kim et al., 2004). Each monomer can be cleaved at a proteasesensitive site into two fragments of 37 kDa and 58 kDa (p37 and p58 moieties) that may represent the A and B moieties of AB-like bacterial toxins. The 37-kDa, amino-terminal portion is highly conserved at the sequence level and is able to induce vacuoles when the vacA gene is placed under the control of a strong eukaryotic promoter and transfected into epithelial cells. This evidence suggests that the active site could be located in this region of the molecule, whereas the carboxy-terminal portion is likely to be devoted to receptor recognition and binding. Although VacA is exported over the outer membrane and is released from the bacteria, recent data have been presented to show that a portion of the toxin remains associated with the bacterial surface. Surface-associated toxin is biologically active and organized into distinct toxin-rich domains on the bacterial surface. Upon bacterial contact with host cells, toxin clusters are transferred to the host cell surface via a contactdependent mechanism, followed by uptake and intoxication (Ilver et al., 2004). The mechanism of toxicity exploited by this virulence factor has not yet been completely elucidated. What is known is that VacA causes an alteration of the endocytic pathway, which results in the selective swelling of late endosomes or prelysosomal structures. The small GTP-binding protein Rab7 is necessary for vacuole formation (Papini et al., 1994, 1997). Even though it is unknown, the target of VacA action is strongly believed to be a fundamental effector in membrane trafficking.

Streptococcus pyogenes NAD+ Glycohydrolase NAD+ glycohydrolase is an important virulence factor produced by group A streptococci (GAS),

which is thought to enhance pathogenicity by facilitating the spread of the microorganism through host tissues. This enzyme catalyzes the hydrolysis of the nicotinamide-ribose bond of NAD to yield nicotinamide and ADP-ribose. Differently from ADP-ribosylating toxins, NAD+ glycohydrolases possess a much higher rate of NADase activity and do not require an ADPribose acceptor. Interestingly this GAS virulence factor is functionally linked to streptolysin O (SLO), a pore-forming toxin, which has been shown to be required for efficient translocation of NAD+ glycohydrolase into epithelial cells. In contrast to the wildtype GAS, isogenic mutants deficient in the expression of SLO, NAD+ glycohydrolase, or both proteins resulted in reduced cytotoxicity and keratinocyte apoptosis. These results suggest that NAD+ glycohydrolase modulates host cell signaling pathways and contributes to the enhancement of streptolysin O cytotoxicity (Bricker et al., 2002).

Toxins Injected into Eukaryotic Cells See Tables 1 and 2 for a summary of the principal features of toxins described in this section. In the classical view, toxins were believed to be molecules that cause intoxication when released by bacteria into the body fluids of multicellular organisms. This definition failed to explain the pathogenicity of many virulent bacteria such as Salmonella, Shigella and Yersinia, which did not release toxic proteins into the culture supernatant. Today we know that these bacteria also intoxicate their hosts by using proteinaceous weapons. These bacteria intoxicate individual eukaryotic cells by using a contact-dependent secretion system to inject or deliver toxic proteins into the cytoplasm of eukaryotic cells (Fig. 2, panel 3). This is done by using specialized secretion systems that in Gramnegative bacteria are called “type III” or “type IV,” depending on whether they use a transmem-

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brane structure similar to flagella or conjugative pili, respectively.

Mediators of Apoptosis Pathogens use different mechanisms to induce or prevent apoptosis in host cells. Virulence factors produced by the pathogen can interact directly with effector molecules of apoptosis or interfere with factors involved in cell survival (Weinrauch and Zychlinsky, 1999). They include: pore-forming toxins which induce cell death by altering host cell permeability, bacterial toxins (such as DT, PAETA, Shiga and Shiga-like toxins) which induce cell death by inhibition of host protein synthesis, and type III secreted proteins of Shigella, Salmonella and Yersinia which are directly delivered into host cell compartment and trigger apoptosis by altering the signal transduction pathway. This latter class of toxins will be described here in more detail.

IpaB Shigella, the causative agent of bacillary dysentery produces IpaB. Shigella invades the epithelial cells by causing the cell cytoskeleton to reorganize during bacterial entry. The bacteria are phagocytosed by macrophages and rapidly escape from phagosomal compartment to the cytosol where they induce apoptosis of the macrophages. Invasion and cytotoxicity require Shigella invasion plasmid antigen (Ipa) proteins, which are secreted by a type III secretion apparatus. Invasion and escape from the phagosome are dependent upon the expression and secretion of the IpaB, IpaC and IpaD. Only IpaB is required to initiate cell death by interaction with the interleukin-1b converting enzyme, or caspase I, which is one of the effector molecules of apoptosis. The IpaB-induced apoptosis results in an inflammation that has the effect not only of clearing and possibly localizing the infection but also promoting bacterial spread in the intestinal epithelium (Hilbi et al., 1998). Protein domains directly involved in pathogenicity have recently been mapped (Guichon et al., 2001).

SipB An analog of Shigella invasin IpaB, Salmonella invasion protein (SipB) is produced by Salmonella and is delivered to the host cells by a type III secretion system. In contrast to Shigella, Salmonella does not escape from the phagosome, but it survives and multiplies within the macrophages. Salmonella virulence genes responsible for invasion and killing of macrophages are encoded by a chromosomal operon named sip

CHAPTER 1.28

containing five genes (sipEBCDA; Hermant et al., 1995). The sip genes show high sequence homology with the ipa operon of Shigella, and the Sip proteins show functional similarities with Ipa proteins. Both proteins have a predominant alpha-helical structure and contain two helical transmembrane domains, which insert deeply into the bilayer (Hume et al., 2003). Similarly to IpaB, SipB also induces apoptosis by binding interleukin-1b-converting enzyme. Necessary for Salmonella-induced macrophage apoptosis, SipB acts through a caspase-Iactivating mechanism similar to that used by IpaB (Hersh et al., 1999). Also, SipB can complement IpaB mutants, enabling them to invade cells and escape macrophage phagosomes.

YopP, YopJ and Related Proteins Yersinia enterocolitica and Yersinia pestis produce YopP and YopJ, respectively (Straley et al., 1986; Mills et al., 1997). Following contact with the host cell, Yersiniae deliver into the cytoplasm of eukaryotic cells, through a type-III secretion system, plasmid-encoded proteins named “Yersinia-outer-membrane proteins” (Yop). These proteins are able to induce alteration of cytoskeleton (YopE and YopT), inhibition of phagocytosis (YopH), and in the case of YopP and YopJ, induction of apoptosis. The mechanism by which Yersinia induces apoptosis is probably different from that described for Shigella, inasmuch as Yersinia induces apoptosis from the outside of host cells. The binding of YopJ directly to the superfamily of MAPKKs blocks both their phosphorylation and subsequent activation. These activities of YopJ are responsible for the inhibition of extracellular signal-regulated kinase, downregulation of TNF-a and suppression of the nuclear factor kappa B (NF-kB) signaling pathways, preventing cytokine synthesis and promoting apoptosis (Orth et al., 1999). The YopJ-related proteins that are found in a number of bacterial pathogens of animals and plants, such as AvrRxv from Xanthomonas campestris (Whalen et al., 1993), AvrA from Salmonella (Hardt et al., 1997), and y410 from Rhizobium (Freiberg et al., 1997) may function to block MAPKKs so that host signaling responses can be modulated upon infection. Whereas no function is known for AvrA and y410, AvrRxv is a plant pathogen virulence protein involved in the programmed cell death pathway.

Toxins Interfering with Inositol Phosphate Metabolism: SopB and IpgD The SopB protein, secreted by Salmonella dublin, is a virulence factor essential for

CHAPTER 1.28

Bacterial Toxins

Motif 1 VVTFNFGVNELALKM SopB IpgD VAAFNVGVNELALKL PTPaseI PVLFNVGINEQQTLA PTPaseII PVLFNVGINEQQTLA

Motif 2 AWNCKSGKDRTGMMSDE CWNCKSGKDRTGMQDAE FTSCKSAKDRTAMSVTL FTCCKSAKDRTSMSVTL

Fig. 45. Alignment of conserved motifs.

enteropathogenicity. The toxin hydrolyzes phosphatidylinositol triphosphate (PIP3), which is a messenger molecule that inhibits chloride secretion, thus favoring fluid accumulation and diarrhea (Norris et al., 1998). Furthermore, SopB, mediates actin cytoskeleton rearrangements and bacterial entry in a Rac-1 and Cdc42-dependent manner. Consistent with an important role for inositol phosphate metabolism in Salmonellainduced cellular responses, a catalytically defective mutant of SopB failed to stimulate actin cytoskeleton rearrangements and bacterial entry (Zhou et al., 2001). SopB is homologous to the Shigella flexneri virulence factor IpgD, suggesting that a similar mechanism of virulence is also present in Shigella. Both proteins contain two regions of sequence similarities (motifs 1 and 2, Fig. 45) with human inositol polyphosphatases types I and II. Motif 2 contains a consensus sequence (Cys-X5-Arg) characteristic of Mg+2-independent phosphatases in which the cysteine is the residue essential for catalysis. Recent studies have shown that IpgD acts as a potent inositol 4phosphatase and is responsible for dramatic morphological changes of the host cell, ultimately leading to consistent actin filament remodeling (Niebuhr et al., 2002).

Toxins Acting on the Cytoskeleton PSEUDOMONAS AERUGINOSA EXOENZYME S. This toxin is one of several products of Pseudomonas aeruginosa that contributes to its pathogenicity (Woods et al., 1989; Kulich et al., 1993; Fig. 1, panel 22). It belongs to the group of ADP-ribosylating factors that lack both the receptor-binding and translocation domains, and are directly injected by bacteria into the cytoplasm of eukaryotic cells. In this case, bacteria intoxicate individual eukaryotic cells by means of a contact-dependent type III secretion system (Yahr et al., 1996). The 49-kDa ExoS protein ADP-ribosylates the small GTP-binding protein Ras at multiple sites but preferably at Arg-41 (Ganesan et al., 1998; see the section ADP-Ribosyltransferases: A Family of Toxins Sharing the Same Enzymatic Activity in this Chapter). To become enzymatically active, ExoS requires the interaction with a cytoplasmic activator named “FAS” or “14.3.3”

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(Fu et al., 1993). When cells are transfected with the exos gene under the control of a eukaryotic cell promoter, a collapse of the cytoskeleton and a change of the morphology of the cells can be observed as primary consequences. Pseudomonas aeruginosa ExoS is a bifunctional cytotoxin where the ADP-ribosyltransferase domain is located within its C-terminus portion. Recent studies showed, in fact, that when transfected or microinjected into eukaryotic cells, the N-terminus part of ExoS (amino acid residues 1–234) stimulates cell rounding. The N-terminus of ExoS (1–234) does not influence nucleotide exchange of Rho, Rac and Cdc42 but increases GTP hydrolysis. It has also been shown that Arg-146 of ExoS is essential for the stimulation of GTPase activity of Rho proteins (Goehring et al., 1999). The GTPase activating domain (GAP) of ExoS has been crystallized (Wurtele et al., 2001). In addition to these toxic effects performed on the cytoskeleton, other activities have been demonstrated for Exo S, such as the adhesive property on buccal cells (Baker et al., 1991) and the induction of human T lymphocyte proliferation (Mody et al., 1995). From sequence analysis, it has been possible to identify the regions of Exo S, which could be involved in NAD binding and thus constitute the common structure of the catalytic site. CLOSTRIDIUM BOTULINUM EXOENZYME C3 AND RELATED PROTEINS. Produced by certain strains of Clostridium botulinum types C and D, exoenzyme C3 is a 251-amino acid protein that specifically ADPribosylates rho and rac gene products in eukaryotic cells (Moriishi et al., 1993; Fig. 1, panel 23). These substrates belong to the group of small GTP-binding proteins and seem to have a fundamental role in cell physiology and cell growth. The ADP-ribosylation process occurs at asparagine residues (Asn-41) located in the putative effector binding domains of rho and rac and thus alter their functions (Sekine et al., 1989). The enzymatic activity is identical to that of all ADPribosylating enzymes; however, the recently solved 3D structure has shown that the C3 exoenzyme structure can be distinguished by the absence of the elongated a-helix, which generally constitutes the ceiling of the active site cleft in the ADP-ribosylating toxins crystallized so far. Seemingly, this feature does not impair the ability of C3 either to accommodate the NAD substrate or to carry out the enzymatic reaction (Han et al., 2001; Fig. 46). This exoenzyme is the prototype of the group of A-only toxins because it apparently lacks the receptor-binding B domain and thus is unable to enter the cells; for this reason, C3 cannot be considered a real virulence factor, and still unknown is whether C3 alone is able to intoxicate the cells.

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Nevertheless, when microinjected into cells, it causes complete disruption of actin-stress fibers, rounding of the cell body, and formation of arborescent extensions. Other members of this family of C3-related exoenzymes have been isolated from Grampositive bacteria, such as certain strains of Staphylococcus aureus (Sugai et al., 1992), Clostridium limosum (Just et al., 1992) and Bacillus cereus (Just et al., 1995c). Whereas C. botulinum C3 and C. limosum exoenzyme are about 70% homologous and immunologically related, the epidermal cell differentiation inhibitor (EDIN) produced by S. aureus is only 35% homologous with C3 and shows no immunological crossreactivity (Fig. 47). However, crystal data recently obtained for S. aureus C3 exotoxin (EDIN-B) have disclosed a very similar structure (Evans et al., 2003). Bacillus cereus exoenzyme exhibits the same substrate specificity as the other C3-like transferases (it was found to act specifically on rho proteins). Nevertheless some differences can be observed for this toxin, such

Fig.46. Crystal structure of exoenzyme C3 of C. botulinum. The residues which constitute the catalytic site are in blue.

CHAPTER 1.28

as the higher molecular weight (28 kDa) and, more importantly, the lack of immunological relationship to any other member of this family (Just et al., 1995a). SALMONELLA SOPE AND SIPA. Salmonella typhimurium achieves entry into cells by delivering effector proteins into the cytosol through a type III secretion system. These effectors stimulate signal pathways leading to reorganization of the cell’s actin cytoskeleton, membrane ruffling and stimulation of nuclear response to promote efficient bacterial internalization. One of the proteins that stimulate the cellular response is SopE, which is able to activate signaling pathways through Rho GTPases by stimulating GTP/GDP nucleotide exchange on proteins such as Cdc42 and Rac (Hardt et al., 1998). These signaling events lead to the recruitment of cellular proteins such as actin and T-plastin (an actin-binding protein that bundles actin), which finally induce actin cytoskeleton rearrangement and membrane ruffling. In addition, SopE stimulates nuclear responses that induce the synthesis of proinflammatory cytokines that contribute to the induction of diarrhea. These cytoskeletal rearrangements are further modulated by SipA, which binds directly to actin, stabilizes actin filaments inhibiting depolymerization, and forms a complex with T-plastin thus increasing its actin-bundling activity (Zhou et al., 1999a, 1999b). SipA activities result in localized actin cytoskeleton reorganization and more pronounced extension of membrane ruffles, which facilitate bacterial uptake. The actin-cytoskeleton reorganization induced by Salmonella is reversible and infected cells are able to recover their normal architecture after bacterial internalization. Crystal structures are available for SipA (Lilic et al., 2003) and for the catalytic fragment of SopE in complex with its host cellular target

Fig. 47. Multiple sequence alignment of protein toxins belonging to the group of exoenzyme C3-like ADP-ribosyltransferases.

CHAPTER 1.28

Bacterial Toxins

Fig. 48. Crystal structures of SipA (panel A) and of SopE in complex with Cdc42 (panel B).

935

Cdc42

SopE

PANEL A

Cdc42 (Buchwald et al., 2002; Figs. 1 [panels 24 and 25] and 48). SHIGELLA IPAA. The entry of Shigella into epithelial cells requires the Ipa proteins, which are secreted upon cell contact by the type III apparatus and act in concert. The IpaB and IpaC proteins form a complex that binds B1 integrin and CD44 receptors and induces actin polymerization at the site of bacterium-cell contact, allowing the formation of membrane extension that probably requires also the action of Cdc42, Rac and Rho GTPases (Nhieu and Sansonetti, 1999). The translocation of IpaA into the cell cytosol probably favors Shigella entry. The IpaA protein binds with high affinity to the N-terminal residues 1–265 of vinculin, a protein involved in linking actin filaments to the plasma membrane. The vinculin-IpaA complex interacts with F-actin inducing subsequent depolymerization of actin filaments. Presumably, these interactions further modulate the formation on the membrane of adhesion-like structures required for efficient invasion. Shigella internalization still occurs at low levels in the absence of IpaA, suggesting that IpaA acts in concert with other bacterial effectors to promote cell entry. Binding of the Shigella protein IpaA to vinculin induces F-actin depolymerization (Bourdet-Sicard et al., 1999). The IpaA and vinculin rapidly associate during bacterial invasion. Although defective for cell entry, an ipaA mutant is still able to induce foci of actin polymerization but differs from wildtype Shigella in its ability to recruit vinculin and a-actinin. It has been postulated that IpaA-vinculin interaction initiates the formation of focal adhesion-like structures required for efficient invasion (Tran Van Nhieu et al., 1997). YERSINIA YOPE. A protein secreted by Yersinia through a type III secretion system,

PANEL B

YopE contributes to the ability of Yersinia to resist phagocytosis (Rosqvist et al., 1990). Following infection of epithelial cells with Yersinia, the microfilament structure of the cells changes leading to a complete disruption of the actin microfilaments, which finally results in cell rounding and detachment from the extracellular matrix (Rosqvist et al., 1991). The effector YopE was recently shown to possess GAP activity towards the Rho GTPases RhoA, Rac and CDC42 in vitro (Aili et al., 2003; Fig. 1, panel 26). Further experimentation has shown that in vivo YopE is able to inhibit Rac- but not Rho- or Cdc42-regulated actin structures. Furthermore, the structure of this toxin has recently been solved, showing a close relationship with the analogous ExoS Gap domain (Evdokimov et al., 2002). YERSINIA YOPT. YopT is the prototype of a new family of 19 cysteine proteases with potent

Fig. 49. Crystal structure of YopE catalytic domain.

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effects on host cells. These include the AVr protein of the plant pathogen Pseudomonas and possibly Yop-J of Yersinia. YopT cleaves the posttranslationally modified cysteine located at the C-terminal end of Rho GTPases (DKGCASS), causing the loss of the prenyl group from RhoA, Rac and cdc42, and releasing them from the membrane (Shao et al., 2003). The inability of Rho to be located to the membrane causes disruption of the cytoskeleton. While the C terminus of YopT is crucial for activity, the N terminus of YopT is crucial for substrate binding (Sorg et al., 2003). SHIGELLA VIRA. The invasiveness of Shigella is an essential pathogenic step and a prerequisite of bacillary dysentery. VirA is a Shigella effector protein, which is delivered into the host cell by a specialized type III secretion system. This protein can interact with tubulin to promote microtubule destabilization and membrane ruffling (Yoshida et al., 2002). With this mechanism, Shigella is able to remodel the cell surface and thus promote its entry into the host. Recent data have shown that VirA deletion mutants displayed decreased invasiveness and were unable to stimulate Rac1.

Toxins Acting on Signal Transduction YERSINIA YPKA AND YOPH. Phosphorylation is central to many regulatory functions associated with the growth and proliferation of eukaryotic cells. Bacteria have learned to interfere with these key functions in several ways. The best-known system is that of Yersinia, where a protein kinase (YpkA; Barz et al., 2000) and a protein tyrosine phosphatase (YopH; Zhang, 1995; Fig. 1, panel 27) are injected into the cytoplasm of eukaryotic cells by a type III secretion system to paralyze the macrophages before they can kill the bacterium. YpkA is a Ser/Thr protein kinase that also displays autophosphorylating activity in vitro. In vivo experiments have shown that this protein is essential for virulence: in fact, challenge with a YpkA knockout mutant causes a nonlethal infection, whereas all mice challenged with wildtype Y. pseudotuberculosis die. Recently, natural eukaryotic substrates of YpkA have been identified by using a two-hybrid assay. These belong to the class of small GTPases and comprise RhoA and Rac-1, but not Cdc42. YopH is a modular protein where the tyrosine phosphatase domain shows a structure and catalytic mechanism very similar to those of eukaryotic enzymes. YopH acts by dephosphorylating cytoskeletal proteins thus disrupting phosphotyrosine-dependent signaling pathways necessary for phagocytosis. Host protein targets include Crk-associated substrate, paxillin, and

CHAPTER 1.28

Fig. 50. X-ray structure of YopH. Colors have been assigned on the basis of secondary structure (yellow for helix and pink for b-sheet). The PTPase phosphate-binding loop and Cys403 are in blue.

focal adhesion kinase. In vivo, YopH inhibits phagocytosis by polymorphonuclear leukocytes (PMNs) and macrophages (Fallman et al., 1995; Ruckdeschel et al., 1996). The protein has a molecular weight of 51 kDa and is composed of an N-terminal domain important for translocation and secretion (Sory et al., 1995) and a Cterminal domain homologous to eukaryotic PTPases (Guan and Dixon, 1990; Bliska, 1995). The three-dimensional structure of YopH has been solved (Stuckey et al., 1994; Su et al., 1994) revealing the presence of a catalytic domain which, despite its low level of sequence identity to the human PTP1B, still contains all of the invariant residues present in eukaryotic PTPases. Its tertiary fold is a highly twisted a/b structure with an eight-stranded b-sheet flanked by seven a-helices. Residues 403–410 form the PTPase phosphate-binding loop with the invariant Cys403 thiol centered within the loop (Fig. 50). EPEC TIR. A 78-kDa protein produced by enteropathogenic E. coli (EPEC) strains, Tir mediates the attachment of bacteria to eukaryotic cells and is essential for EPEC virulence. The Tir protein is tyrosine phosphorylated upon injection into eukaryotic cells by a type III secretion system. While in the host cell, it becomes an integral part of the eukaryotic cell membrane and functions as receptor for intimin, the major EPEC adhesin (Kenny et al., 1997). It is believed that, once in the host, Tir adopts a hairpin-like structure using its two putative transmembrane domains (TMDs) to span the host cell membrane. The region between the two TMDs constitutes the extracellular loop that functions as the intimin-binding domain. Following tyrosine

CHAPTER 1.28

phosphorylation, the protein mediates actin nucleation, resulting in pedestal formation and triggering tyrosine phosphorylation of additional host proteins, including phospholipase C-g. Tir is essential for EPEC virulence and was the first bacterial protein described to be tyrosine phosphorylated by host cells (Crawford and Kaper, 2002). HELICOBACTER PYLORI CAGA. Cytotoxin-associated gene A (CagA) is an immunodominant protein produced by most virulent strains of Helicobacter pylori, with a size that can vary from 128 kDa to 146 kDa and which is commonly expressed in peptic ulcer disease (Covacci et al., 1993b). CagA is characterized by a central region containing an EPIYA motif, which can be repeated up to six times increasing the molecular weight of the protein. The gene is encoded within a pathogenicity island, which also encodes the type IV secretion system necessary to inject the protein into eukaryotic cells. Once injected into the host cell, the protein is tyrosine phosphorylated at the EPIYA motif by the kinase C-Src and Lyn. The signal is proportional to the number of EPIYA motives present (Stein et al., 2000). The tyrosine phosphorylated CagA (CagA-P) activates SHP-2, inactivates C-Src leading to cortactin dephosphorylation triggering a signal transduction cascade (which results in cellular scattering proliferation, a phenotype indistinguishable from that induced by the hepatocyte growth factor [HGF]). The long-term chronic infection and the continuous stimulation increase the risk of cancer of people infected by CagA+ H. pylori. CagA is the first bacterial protein linked to cancer in humans and the cagA gene can be considered the first bacterial oncogene. YERSINIA PESTIS YOPM. YopM is an effector protein delivered to the cytoplasm of infected cells by the type III secretion mechanism of Yersinia pestis. YopM is a highly acidic protein, which is essential for virulence, but whose mechanism of action is still elusive. Differently from other effectors, this toxin has been shown to accumulate not only in the cytoplasm but also in the nucleus of mammalian cells. Recently, McDonald and colleagues have found that YopM interacts with two kinases, protein kinase C-like 2 (PRK2) and ribosomal S6 protein kinase 1 (RSK1). These two kinases associate only when YopM is present, and expression of YopM in cells stimulates the activity of both kinases. These results indicate that PRK2 and RSK1 are the first intracellular targets of YopM (McDonald et al., 2003). The X-ray structure determined for YopM has shown a modular architecture constituted by leucine-rich repeats, mainly organized in an extended b-sheet structure (Evdokimov et al.,

Bacterial Toxins

937

Fig. 51. Crystal structure of YopM effector protein of Yersinia pestis.

2001; Figs. 1 [panel 28] and 51). This organization is very similar to that found for other important proteins, such as rab geranylgeranyltransferase and internalin B produced by Listeria. SALMONELLA SPTP. Salmonella protein tyrosine phosphatase (SptP) is an effector protein secreted by the type III secretion apparatus of Salmonella enterica. SptP is a modular protein composed of two functional domains, a C-terminal region with sequence similarity to Yersinia tyrosine phosphatase YopH, and an N-terminal domain showing homology to bacterial cytotoxins such as Yersinia YopE and Pseudomonas ExoS (Murli et al., 2001). Recently, it was demonstrated that this domain possesses strong GTPase activating domain protein (GAP) activity for Cdc42 and Rac1. The crystal structure of SptP-Rac1 complex has shown that SptP is strongly stabilized by this interaction (Stebbins and Galan, 2000; Fig. 52). PSEUDOMONAS AERUGINOSA EXOU. Several extracellular products secreted by the P. aeruginosa type III secretion system are responsible for virulence. Among these, the 70-kDa protein, ExoU, is responsible for causing acute cytotoxicity in vitro and epithelial lung injury. Recent studies demonstrated that ExoU has lipase activity, and that the cytotoxicity of ExoU is dependent on its patatin-like phospholipase domain. The results suggest that ExoU requires the presence of a catalytically active site Ser(142) and that a yet unknown eukaryotic cell factor(s) is necessary for its activation (Tamura et al., 2004).

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CHAPTER 1.28 Fig. 52. Crystal structure of SptP in complex with Rac-1 (yellow). The Nterm and C-term domains of SptP are colored in cyano and purple, respectively.

Rac-1 Ct Nt

NH2

N

N

Target protein

CO3NH3

+

O N

N

Target protein O

OH OH

N

N

N

O

H C O F O P O CH3 O 3 O OH OH OH OH

NH2 N

O

N

O

H3C O F O P O CH3 OH OH

OH OH

O

CO3NH3

+

N

OH OH

(Cys, Arg, Asn, Diphinarmide)

NAD

ADP-ribose

nicotinamide

Fig. 53. Mechanism of ADP-ribosylation reaction catalyzed by ADP-ribosyltransferases.

ADP-Ribosyltransferases: A Family of Toxins Sharing the Same Enzymatic Activity ADP-Ribosylating Toxins: Main Features The ADP-ribosylating toxins are a class of bacterial proteins that characterized by an enzymatic domain with ADP-ribosyltransferase activity (Ueda and Hayaishi, 1985; Althaus and Richter, 1987). During ADP-ribosylation (Fig. 53), these toxins bind NAD and transfer the ADP-ribose moiety to a specific substrate molecule, which is thus forced to undergo a dramatic functional modification. The toxic effect is totally dependent upon the enzymatic activity. On the basis of their overall structure, ADPribosyltransferases can be separated into A/B toxins, binary toxins, and A-only toxins, where A is the subunit with the enzymatic activity, and B is the carrier domain involved in the recognition of the specific surface receptor and in the translocation of the toxic moiety into the eukaryotic cell. Most of the best characterized ADPribosylating toxins belong to the class with an A/B architecture: pertussis toxin (PT; Locht et al., 1986; Nicosia et al., 1986), cholera toxin (CT; Mekalanos et al., 1983), and E. coli heat-labile enterotoxin (LT; Spicer and Noble, 1982; Yamamoto et al., 1984) are typical examples of this

family where the A domain (called “S1” in PT) bears the enzymatic core and the B domain is an oligomer that helps the translocation across the cell membrane; the two subunits are linked together by noncovalent bonds. The genes coding for CT and LT are highly homologous (Dallas and Falkow, 1980) and are organized into operons located on the chromosome of Vibrio cholerae and on a plasmid of E. coli (So et al., 1978). Diphtheria toxin (DT; Pappenheimer, 1977; Collier et al., 1982) and Pseudomonas aeruginosa exotoxin A (PAETA; Gray et al., 1984b; Wick et al., 1990) are A/B toxins with a three-domain structure: the catalytic domain C, contained in fragment A, and the transmembrane domain T and receptor-binding domain R, both within the B subunit. The binary (as opposed to the A/B) toxins have a fairly similar organization, but in this case the A and B domains are separately secreted in the culture supernatant where the B domain initially binds the receptor on the surface of the target cell and only then is able to bind the A subunit and help its translocation into the cytosol. Examples of this family of ADP-ribosyltransferases are the C2 toxin of Clostridium botulinum (Aktories et al., 1986), the iota toxin of C. perfringens (Perelle et al, 1995), the toxin of C. spiroforme (Popoff and Boquet, 1988a), the mosquitocidal toxin (MTX) of Bacillus sphaeri-

CHAPTER 1.28

Bacterial Toxins

cus (Thanabalu et al., 1993), and the C. difficile transferase (Just et al., 1994). Finally, the “A-only” toxins include Exo S of Pseudomonas aeruginosa (Kulich et al., 1994) and other toxins such as C3 of Clostridium botulinum (Nemoto et al., 1991), EDIN of Staphylococcus aureus (Sugai et al., 1990), and the toxins of Bacillus cereus (Just et al., 1995b) and of Clostridium limosum (Just et al., 1992). All the A-only toxins possess a still unknown mechanism of cell entry, with the notable exception of Exo S, which has been shown to be directly injected into eukaryotic cells by a specialized secretion system (Yahr et al., 1996). With the exception of actin, all the eukaryotic proteins that are ADP-ribosylated by these toxins are GTP-binding proteins (G-proteins); these proteins are molecular switches involved in a number of essential cell functions including protein synthesis and translocation, signal transduction, cell proliferation, and vesicular trafficking (Hamm and Gilchrist, 1996).

ADP-Ribosylating Toxins: A Common Structure of the Catalytic Site Bacterial enzymes with ADP-ribosyltransferase activity include a variety of toxins with different

939

structural organizations; the better-represented class is that comprising proteins with an A/B structure (PAETA, DT, CT, LT and PT), where subunit A is responsible for enzymatic activity and subunit B is involved in receptor binding. Other toxins, termed “binary toxins” (Clostridium botulinum toxin C2 and related proteins) are still composed of the two functional domains A and B. However, they reside on different molecules and need to interact to acquire activity. Finally, there is a group of ADPribosylating toxins that do not possess the receptor-binding domain B at all and are thus named “A-only toxins.” This group includes Clostridium botulinum exoenzyme C3 and related proteins, which are unable to invade the cells, and toxins which are directly injected into eukaryotic cells (ExoS) by means of a specialized secretion apparatus. From primary sequence analysis, it is possible to identify two main groups of homology (Fig. 54): the DT-like group, mainly composed of DT and PAETA, and the CT-like group comprising the remaining ADP-ribosyltransferases. Although some homology is present among the members of the CT group, no overall significant and extended sequence similarity can be detected to justify the observed common mech-

CT group Region 2

Region 1

CT LT LT-II PT Exos C2_bot C3_bot C3_II C3_lim IOTA_tox

B.cereus Edin

8

Region 3

IY R A 5 IY R A

55

H D D G Y V S T S I S L R S A H L V G Q T I 76

55

Y D D G Y V S T S L S L R S A H L A G Q S I 76

4

50

Y N D G Y V S T T V T L R Q A H L I G Q N I 74 F V S T S S S R R Y T E V Y L E H R M Q E 76

FFR A 10 VYR A

... 320 AYR A LFR G IFR G LFR G V Y R R 287 ... 121 VYR L

230

D D G Y L S T S L N P G V A R S G Q G T I 150 345

S F STSL K S T P LS F S G Y I S T S L M S A Q FG G R

131 131

150 149

S T S L M N V S Q F A G 149 S T S L V N G S A F A G R 147 S T S I G S V N M S A F A K R K I 154 ...... Y S S T Q L V S G A A V G G R 100

G YI 131 G YI 136 FI 178

G

E Q E V S A L 316 E Q E V S A L 316 113 106 Y P S E N EF A A L 176 T Y Q S E Y L 110 378 Y K N E K E L L Y 383 305 I E Q E I L L N 383 169 F P Q Q L E V L L P 178 169 F A Q Q L E M L L P 178 169 F K Q Q L E V L L P 178 373 Y A G E Y E V L L 303 ... Y P Q Q Y E L L L P ...... 219 210 Y YQ QQEV L LP 107 107

HPD HPY

DT group D T

S Y H G T 23

PAETA

G Y H G T 443

50

W K G F YS T D N K Y D A A GY

15

14

444

W R G F YI A G D P A L A Y GY

101

55

E E

YI N TI L

Fig. 54. Sequence alignment of protein segments containing Regions 1, 2 and 3 of bacterial ADP-ribosylating enzymes. The two groups of homology (DT-like and CT-like groups) are distinguished. Catalytic residues of Regions 1 and 3, and most relevant and conserved residues of Region 2 are colored in red; extended consensus sequences detected in the three regions are boxed, whereas other partially conserved residues are in boldface. Predicted and observed secondary structure folding is indicated for each region: Regions 1 and 3 are b-strands (arrows), while Region 2 is characterized by a short coil (solid line), followed by a b-strand and by an a-helix.

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anism of catalysis; nevertheless, biochemical experiments of photoaffinity labeling and studies of site-directed mutagenesis had previously demonstrated for most of the toxins that the presence of a glutamic acid is so important for catalytic activity, even a conservative substitution with an aspartate could not be tolerated without loss or drastic decrease of toxicity (Douglas and Collier, 1987; Wilson et al., 1990; Lobet et al., 1991; Antoine et al., 1993). On the basis of these experimental data and on the crystallographic structures which are now available for LT (Sixma et al., 1991), CT (Zhang et al., 1995), PT (Stein et al., 1994), DT (Choe et al., 1992) and PAETA (Allured et al., 1986), a common catalytic site could be identified which, despite the low level of sequence homology, is almost perfectly superimposable for all them (Domenighini et al., 1994). In terms of tertiary structure, the active site is a cleft formed by a b-strand followed by a slanted a-helix that has a different length in the various toxins (spanning from 12 residues for DT, PAETA and LT, to 21 in the case of PT). The bstrand and the a-helix represent, respectively, the lower and upper face of the cavity in which the nicotinamide ring of NAD is anchored during the enzymatic reaction (Region 2 of Fig. 54). Although all the toxins share this similar folding in the region of the active site, at the amino acid level, the only residue which is well conserved among all the representatives of the CTand DT-groups is a glutamic acid (Glu-148 of DT, Glu-553 of PAETA, Glu-112 of CT and LT, and Glu-129 of PT), which corresponds to the core of Region 3 (Fig. 54). These residues retain an equivalent spatial position and orientation residing in a short b-strand flanking the external side of the cavity (Fig. 55). With the exception of the conserved glutamate, the consensus sequence generated for Region 3 differs between the two groups of toxins. In the DT family, in fact, it is composed of the catalytic Glu followed by an aromatic and a hydrophobic residue, whereas in the CT-group, the consensus can be extended to a few neighboring residues (Fig. 45). On the basis of alignment of C2-I with iota toxin and with the other ADP-ribosyltransferases, the catalytic glutamate was identified (Glu-389 of C2) and its function experimentally confirmed by sitedirected mutagenesis (Barth et al., 1998). In the case of Pseudomonas aeruginosa Exo S, the equivalent Glu has been mapped at position 381 (Liu et al., 1996). Another well-conserved residue is His-21 of DT that can be aligned to His-440 of PAETA, and with the conserved Arg-7 of CT and LT, and Arg-9 of PT (Burnette et al., 1988, 1991; Papini et al., 1990; Lobet et al., 1991; Han and Galloway, 1995). The segment comprising this residue is

CHAPTER 1.28

Region 2 PT LT Tyr65 Glu

Region 3

DT

PAETA

Region 1 Tyr54

Arg/His

Fig. 55. Superimposition of the three-dimensional structures of the NAD-binding cavities (Region 2) of the bacterial toxins LT (green), PT (yellow), DT (red) and PAETA (blue). The catalytic residues carried by Region 1 (Arg/His) and by Region 3 (Glu) and common to the two regions of homology are shown. In addition, the two essential tyrosines of the DT group are colored in red.

termed “Region 1” (Fig. 54). These amino acids are once again located in essentially identical positions within the active site, lying opposite to the glutamic acid on an antiparallel b-strand close to the internal face of the catalytic cleft. Although several models have been proposed to explain the possible function of the conserved histidine/arginine of Region 1, that this residue does not play a direct role in catalysis seems now widely accepted; very likely it may have a function in maintaining the integrity of the active-site pocket upon formation of structurally stabilizing hydrogen bonds (Johnson and Nicholls, 1994). Nevertheless, mutations at the His-440 position of PAETA, though affecting the enzymatic activity, have little or no effect on NAD-binding (Han and Galloway, 1995); this suggests that His-440 may not be exactly homologous to His-21 of DT or to the arginines of the CT-group. In the case of the C2-I component of clostridial toxin C2, site-directed mutagenesis of Arg-299 induced a dramatic reduction of transferase activity, thus suggesting an equivalent role for this residue in the conformation of the active site (Perelle et al., 1995). Region 2 includes a number of amino acids that, while maintaining the same secondary structure in both DT- and CT-families (Fig. 55), result in a major sequence difference (Fig. 54). This is mainly a structural region corresponding to the core of the active site cleft, which is devoted to the docking of NAD. The consensus sequence generated for the DT group is characterized by two conserved tyrosines spaced by ten amino acids, and located on the middle portion

CHAPTER 1.28

of the b-strand and on the internal face of the a-helix, respectively. Tyr-54 and Tyr-65 of DT, and Tyr-470 and Tyr-481 of PAETA have been shown to play an important role in catalysis inasmuch as they anchor the nicotinamide ring during the reaction by creating a p pile of three aromatic rings which strengthen the overall binding of NAD and stabilize the complex (Carroll and Collier, 1984; Li et al., 1995). This consensus motif can be extended to four other residues which precede the first Tyr, and to a glycine residue which is located upstream of the second Tyr. In PT, a similar role is likely to be played by Tyr-59 and Tyr-63, which have a similar spatial orientation and distance from each other. This observation is supported by the fact that in CT and LT, where the stacking interactions produced by the two tyrosines are lacking, the affinity for NAD is 1000-fold lower (Galloway and van Heyningen, 1987). In the case of the CT-group, Region 2 is centered on a consensus core domain characterized by the motif Ser-Thr-Ser that is observed and predicted to fold in a b-strand representing the floor of the cavity. Experiments of site-directed mutagenesis have confirmed the importance of these residues in maintaining the shape of the cavity. Substitutions of Ser-61 and Ser-63 of LT with Phe and Lys, respectively, have been shown to produce nontoxic mutants (Harford et al., 1989; Fontana et al., 1995). The core sequence of Region 2 can be extended to give the more general consensus aromatic-hydrophobic-Ser-ThrSer-hydrophobic. Another amino acid that has been proposed as being important in catalysis is His-35 of PT (Xu et al., 1994) located near the beginning of the bstrand which forms the floor of the cavity, in a position equivalent to that of His-44 of LT and CT (Yamashita et al., 1991); a functional homologue, His is also present in the mosquitocidal toxin SSII-1 from Bacillus sphaericus (Thanabalu et al., 1991) but is absent in DT and PAETA. In the 3D structure, this residue appears to be sufficiently close to the oxygen atom of the ribose ring of NAD to interact with it and increase the electrophilicity of the adjacent anomeric carbon atom. The absence of an equivalent residue in DT and PAETA again supports the idea that the two groups of toxins perform the same enzymatic activity in a slightly different fashion. An additional feature that is common to all ADP-ribosylating toxins is the need for a conformational rearrangement to achieve enzymatic activity. In the native structure, in fact, the NADbinding site of LT and CT is obstructed by a loop (amino acids 47–56) that needs to be displaced to obtain a functional NAD-binding cavity. A

Bacterial Toxins

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functionally homologous region is also present in PT where the loop comprises residues 199–207. In the case of DT, where the crystallographic data of the complex are available, the observation that the active-site loop consisting of amino acids 39–46 changes structure upon NADbinding, suggests that these residues may be important for the recognition of the ADP-ribose acceptor substrate, EF-2 (Weiss et al., 1995; Bell and Eisenberg, 1996). The recent publication of the crystallographic data of the DT-NAD complex, and the presence of common features within all ADP-ribosylating toxins, permits speculation on a possible common mechanism of catalysis (Fig. 56). The best hypothesis is that NAD enters the cavity, which is then made available for the recognition of the

TOXIN

Arg/His Glu NAD

SH

NH2

+

C NH Cysteine

N

NH2 R

Arginine

N H

Diphthamide

SUBSTRATE

Fig. 56. Schematic representation of a possible common mechanism of catalysis: the nicotinamide adenine dinucleotide (NAD) molecule (red) is docked inside the cavity by means of stacking interactions provided by the two aromatic rings (yellow) that protrude from the scaffold of Region 2. The catalytic glutamic acid (purple) and its possible interactions with the acceptor residues of the various substrates are also reported. The Arg/His residue (green) provides stabilizing interactions with the backbone of the cavity and seems to be also responsible for the correct positioning of NAD inside the pocket.

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substrate, upon displacement of the mobile loop. Then, NAD docks at the bottom of the pocket where a small residue (the conserved serine in Region 2 of the CT-group, the threonine-56 of DT, and the alanine-472 of PAETA) is required to allow good positioning. The nicotinamide moiety of NAD is then blocked in a suitable position by means of stacking interactions provided by a couple of aromatic rings (Tyr-54 and Tyr-65 of DT, Tyr-470 and Tyr-481 of PAETA, and possibly, Tyr-59 and Tyr-63 of PT). In this context the conserved arginine/histidine might display its key role in maintaining the correct shape of the active site pocket via hydrogen bonds formed with the backbone of the structure and possibly one with the ribose moiety. The enzymatic reaction is then catalyzed by the essential glutamic acid, which is likely to stabilize a positively charged oxocarbonium intermediate of NAD, to favor its subsequent interaction with the nucleophilic residue of the incoming substrate (diphthamide in the case of DT and PAETA, arginine in the case of LT and CT, and cysteine in the case of PT).

Novel ADP-Ribosylating Toxins Detected by Genome-Mining With the advent of the Genomic Era, identification of bacterial factors possibly involved in virulence is an easier challenge. In fact, given the vast amount of information that we now possess on toxins—including sequence data—and thanks to the growing number of sequenced bacterial genomes, it is possible to proceed by homology criteria to predict novel members of important classes of bacterial toxins. Several examples exist where computer-based methodologies have been instrumental to the identification of novel potential bacterial toxins in sequenced genomes. Among them, we will mention here the case of mono ADPribosyltransferases. Mono-ADP-ribosyltransferases (mADPRTs) constitute a class of potent toxins in bacteria, which generally play an important role in the pathogenesis of related microorganisms. Despite the poor overall conservation at the primary structure level, the catalytic subunits of these toxins show a remarkable similarity within the enzymatic cavity, so that these portions of the proteins are quite well conserved. For these reasons, and encouraged by the availability of a growing number of sequenced bacterial genomes, a series of studies have been directed towards the computer-based identification of novel members of this family of enzymes by means of sequence-homology criteria in finished and unfinished genome sequences. As a result, more than twenty novel putative ADP-

CHAPTER 1.28

ribosyltransferases have been identified both in Gram-positive and Gram-negative organisms, including five from Pseudomonas syringae, five from Burkholderia cepacea, two from Enterococcus faecalis, and one each from Salmonella typhi, Streptococcus pyogenes, Mycoplasma pneumoniae, Streptomyces coelicolor, Bacillus halodurans and Vibrio parahaemolyticus (Pallen et al., 2001). With the exception of the protein detected in Salmonella, which is adjacent to an ORF protein similar to the S2 subunit of pertussis toxin, all the other genome-derived putative ADPRTs lack a predicted translocation domain. So far, none of these bacterial proteins has been tested either for their ADP-ribosyltransferase activity or for the capability of entering eukaryotic cells; however, sequence data indicate a possible role of these proteins in the pathogenesis of the corresponding microorganisms. Very recently, a new protein has been added to the list of ADPribosyltransferases detected by computer analysis (Masignani et al., 2003). This novel factor has been identified by means of primary and secondary structure analysis in the genomic sequence of a virulent isolate of Neisseria meningitidis and has been named “NarE” (Neisseria ADPribosylating enzyme). As predicted by “in silico” studies, biochemical analysis has demonstrated that NarE is capable of transferring an ADPribose moiety to a synthetic substrate.

Toxins with Unknown Mechanism of Action See Tables 1 and 2 for a summary of the principal features of toxins described in this section. The zonula occludens toxin (Zot) is produced by bacteriophages present in toxinogenic strains of Vibrio cholerae. Zot is a single polypeptide chain of 44.8 kDa, which localizes in the outer membranes. After internal cleavage, a carboxyterminal fragment of 12 kDa is excreted and this is probably responsible for the biologic effect. Zot has the ability to reversibly alter the tight junctions of intestinal epithelium, thus facilitating the passage of macromolecules through mucosal barriers (Di Pierro et al., 2001). Zot has also been shown to act as mucosal adjuvant and to induce protective immune response in the animal model (Marinaro et al., 2003). Hemolysin BL (HBL) is an enterotoxin produced by B. cereus, which is composed of three proteins (B, L1 and L2), each with a molecular mass of 40 kDa, and whose corresponding genes are located on the same operon. HBL has hemolytic as well as dermonecrotic and vascular permeability activities and is able to cause fluid accumulation in ligated rabbit ileal loops (Beecher et al., 1997; Beecher and Wong, 2000).

CHAPTER 1.28

The bile-salt hydrolase (BSH) is a protein elaborated by Listeria monocytogenes, which is absent from the genome of the nonpathogenic L. innocua. The bsh gene encodes an intracellular enzyme and is positively regulated by PrfA, the transcriptional activator of known L. monocytogenes virulence genes (Dussurget et al., 2002). Furthermore, bsh deletion mutants show reduced virulence and liver colonization, thus demonstrating that BSH is a toxin specifically involved in the intestinal and hepatic phases of listeriosis.

Literature Cited Abrahmsen, L., M. Dohlsten, S. Segren, P. Bjork, E. Jonsson, and T. Kalland. 1995. Characterization of two distinct MHC class II binding sites in the superantigen staphylococcal enterotoxin A. EMBO J. 14(13):2978–2986. Aili, M., M. Telepnev, B. Hallberg, H. Wolf-Watz, and R. Rosqvist. 2003. In vitro GAP activity towards RhoA, Rac1 and Cdc42 is not a prerequisite for YopE induced HeLa cell cytotoxicity. Microb Pathog 34(6):297–308. Aktories, K., M. Barmann, I. Ohishi, S. Tsuyama, K. H. Jakobs, and E. Habermann. 1986. Botulinum C2 toxin ADP-ribosylates actin. Nature 322:390–392. Aktories, K. 1994. Clostridial ADP-ribosylating toxins: effects on ATP and GTP-binding proteins. Molec. Cell Biochem. 138:167–176. Allured, V. S., R. J. Collier, S. F. Carroll, and D. B. McKay. 1986. Structure of exotoxin A of Pseudomonas aeruginosa at 3.0-Angstrom resolution. Proc. Natl. Acad. Sci. USA 83(5):1320–1324. Alouf, J. E., and C. Geoffrey. 1991. Sourcebook of bacterial protein toxins. In: Alouf and Freer (Eds.) London, UK. 147–186. Althaus, F. R., and C. Richter. 1987. ADP-ribosylation of proteins: Enzymology and biological significance. Molec. Biol. Biochem. Biophys. 37:1–237. Altwegg, M., and H. K. Geiss. 1989. Aeromonas as a human pathogen. Crit. Rev. Microbiol. 16:253–258. Antoine, R., A. Tallett, S. van Heyningen, and C. Locht. 1993. Evidence for a catalytic role of glutamic acid 129 in the NAD-glycohydrolase activity of the pertussis toxin S1 subunit. J. Biol. Chem. 268:24149–24155. Atassi, M. Z., and M. Oshima. 1999. Sructure, activity, and immune (T and B cell) recognition of botulinum neurotoxins. Crit. Rev. Immunol. 19:219–260. Baker, N. R., V. Minor, C. Deal, M. S. Shahrabadi, D. A. Simpson, and D. E. Woods. 1991. Pseudomonas aeruginosa exoenzyme S is an adhesin. Infect. Immunol. 59:2859–2863. Baldwin, R. L., M. S. Kobrin, T. Tran, I. Pastan, and M. Korc. 1996. Cytotoxic effects of TGF-alpha-Pseudomonas exotoxin A fusion protein in human pancreatic carcinoma cells. Pancreas 13:16–21. Ballard, J., J. Crabtree, B. A. Roe, and R. K. Tweten. 1995. epticum alpha-toxin exhibits similarity with that of Aeromonas hydrophila aerolysin. Infect. Immunol. 63:340–344. Barry, E. M., A. A. Weiss, I. E. Ehrmann, M. C. Gray, E. L. Hewlett, and M. S. Goodwin. 1991. Bordetella pertussis adenylate cyclase toxin and hemolytic activities require

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a second gene, cyaC, for activation. J. Bacteriol. 173:720– 726. Barth, H., J. C. Preiss, F. Hofmann, and K. Aktories. 1998. Characterization of the catalytic site of the ADPribosyltransferase Clostridium botulinum C2 toxin by site-directed mutagenesis. J. Biol. Chem. 273:29506– 29511. Barz, C., T. N. Abahji, K. Trulzsch, and J. Heesemann. 2000. The Yersinia Ser/Thr protein kinase YpkA/YopO directly interacts with the small GTPases RhoA and Rac-1. FEBS Lett 29 482(1–2):139–143. Beecher, D. J., and A. C. Wong. 1997. Tripartite hemolysin BL from Bacillus cereus. Hemolytic analysis of component interactions and a model for its characteristic paradoxical zone phenomenon. J Biol Chem 272(1):233– 239. Beecher, D. J., and A. C. Wong. 2000. Tripartite haemolysin BL: isolation and characterization of two distinct homologous sets of components from a single Bacillus cereus isolate. Microbiology 146(Pt 6):1371–1380. Bell, C. E., and D. Eisenberg. 1996. Crystal structure of diphtheria toxin bound to nicotinamide adenine dinucleotide. Biochemistry 35:1137–1149. Bell, C. E., and D. Eisenberg. 1997. Crystal structure of diphtheria toxin bound to nicotinamide adenine dinucleotide. Adv. Exp. Med. Biol. 419:35–43. Bennett, M. J., and D. Eisenberg. 1994. Refined structure of monomeric diphtheria toxin at 2.3 Å resolution. Protein Sci. 3:1464–1475. Bhakdi, S., F. Grimminger, N. Suttorp, D. Walmrath, and W. Seeger. 1994. Proteinaceous bacterial toxins and pathogenesis of sepsis syndrome and septic shock: The unknown connection. Med. Microbiol. Immunol. (Berl.) 183:119–144. Billington, S. J., B. H. Jost, and J. G. Songer. 2000. Thiolactivated cytolysins: structure, function and role in pathogenesis. FEMS Microbiol. Lett. 182:197–205. Blackman, M. A., and D. L. Woodland. 1995. In vivo effects of superantigens. Life Sci. 57:1717–1735. Blanke, S. R., K. Huang, B. A. Wilson, E. Papini, A. Covacci, and R. J. Collier. 1994. Active-site mutations of the diphtheria toxin catalytic domain: role of histidine-21 in nicotinamide adenine dinucleotide binding and ADPribosylation of elongation factor 2. Biochemistry 33:5155–5161. Bliska, J. B. 1995. Crystal structure of the Yersinia tyrosine phosphatase. Trends Microbiol. 3:125–127. Bohach, G. A., D. J. Fast, R. D. Nelson, and P. M. Schlievert. 1990. Staphylococcal and streptococcal pyrogenic toxins involved in toxic shock syndrome and related illnesses. Crit. Rev. Microbiol. 17(4):251–272. Booth, B. A., M. Boesman-Finkelstein, and R. A. Finkelstein. 1984. Vibrio cholerae hemagglutinin/protease nicks cholera enterotoxin. Infect. Immunol. 45:558–560. Bourdet-Sicard, R., M. Rudiger, B. M. Jockusch, P. Gounon, P. J. Sansonetti, and G. T. Nhieu. 1999. Binding of the Shigella protein IpaA to vinculin induces F-actin depolymerization. EMBO J. 18:5853–5862. Braun, V., and T. Focareta. 1991. Pore-forming bacterial protein hemolysins (cytolysins). Crit. Rev. Microbiol. 18:115–158. Brossier, F., M. Weber-Levy, M. Mock, and J. C. Sirard. 2000. Role of toxin functional domains in anthrax pathogenesis. Infect. Immunol. 68:1781–1786. Brown, B. A., and J. W. Bodley. 1979. Primary structure at the site in beef and wheat elongation factor 2 of ADP-

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Prokaryotes (2006) 2:956–968 DOI: 10.1007/0-387-30742-7_29

CHAPTER 1.29 ehT

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The Metabolic Pathways of Biodegradation LAWRENCE P. WACKETT

Introduction History The decay (biodegradation) of organic matter has been a part of life throughout human history. When the organic matter was a person’s food, clothing or dwelling, biodegradation was no doubt very undesirable. In this context, humans have, through most of their history, sought to prevent or, more practically, slow down biodegradation. Animal hides were treated with tannins to crosslink proteins and prevent their degradation. Food was dried, salted or pickled to prevent microbial growth, and hence, spoilage. Though solutions to biodegradation were found, the underlying causes of the phenomenon were less clear. But no doubt, correlations were made by direct observation of macroscopic microorganisms, typically fungi, on rotting material. People could see wood rot fungi on decaying wood or hyphal masses on bread. And, in fact, some of the foundations for the science of microbiology were established with macroscopic fungi. Micheli showed that fungi now known as Mucor, Botrytis and Aspergillis could be cultivated on the surfaces of fresh-cut melon, quince and pear (Bull and Slater, 1982). Micheli serially transferred the fungi, thus initiating the practice of isolating, maintaining and characterizing specific genera of microorganisms. He also observed that different fungal genera showed preferences for certain fruits used as the cultivating medium, thus establishing the idea of selective culture. In fact, these observations became entwined with the controversy over spontaneous generation because macroscopic fungal growth derived from microscopic fungal spores. Many interpreted proliferation of microscopic life, biodegrading different organic material, as the spontaneous generation of life from non-life. Louis Pasteur is generally credited with demonstrating most convincingly that the elimination of all bacterial and fungal contamination would prevent spoilage (Clarke, 1985). It was recognized from the work of Pasteur, Tyndall and

others that bacteria are ubiquitous and difficult to remove from any given environment. In a broad sense, virtually all prokaryotes participate in biodegradation. Prokaryotes decompose (biodegrade) organic molecules as part of their need to derive chemical energy to make ATP or to produce metabolic intermediates. In the early twentieth century, Beijerinck (1901) and Winogradsky (1890) contributed to the current idea that prokaryotes are important in the recycling of carbon, nitrogen, sulfur and other elements on a global scale. For example, we now know that 1015 grams of methane gas are produced annually by anaerobic Archaea known as “methanogens” and most of the biogenic methane is oxidized by aerobic methanotrophic bacteria (Lipscomb, 1994). This constitutes one small part of the global carbon cycle. If one considers that over ten million organic compounds are known, many of which are theoretically biodegradable, the magnitude of these cycles is enormous.

Scope of Biodegradation in the Modern World Although naturally occurring compounds biodegrade on a massive scale, the biodegradation of synthetic compounds attracts more interest. Over the last century, some synthetic, industrial chemicals have been shown to exert toxic or carcinogenic effects on humans. For example, factory workers in aniline dye (Bulbulyan et al., 1995) and vinyl chloride polymer industries (Langard et al., 2000) were developing certain cancers at relatively high rates and the epidemiological studies were confirmed in animals. From these observations has emerged the awareness that, with increasing world population, more effort must be expended to maintain clean water, air and soil. Problems of human exposure to potentially harmful organic compounds can be handled in different ways. For example, certain chemicals may be banned, manufacturing processes can be made cleaner, or wastes can be treated at the

CHAPTER 1.29

source or in the open environment. In fact, all of those are occurring. And prokaryotes are increasingly being exploited for treating wastes, either in the manufacturing facility or for remediation of chemical spills or releases. Some high profile applications have been published (Harkness et al., 1993; Roberts et al., 1993; Strong et al., 2000; Wagner-Dobler et al., 2000) but in fact, most of these applications are quietly in use at manufacturing facilities all around the globe.

The University of Minnesota Biocatalysis/Biodegradation Database To facilitate the use of microbial catalysis, either for developing cleaner manufacturing or treating wastes, we have developed the University of Minnesota Biocatalysis/Biodegradation Database (UM-BBD) (umbbd.ahc.umn.edu). The UM-BBD provides information on microbial biocatalytic non-intermediary metabolism (i.e., reactions generally associated with biodegradation of synthetic industrial chemicals; Ellis et al., 2000). This metabolism is considered nonintermediary because it is restricted to only a few prokaryotes. It is these non-intermediary metabolic reactions that biotransform compounds and funnel them into the central metabolism of most prokaryotes. For example, only a limited number of prokaryotes can catabolize nitrobenzene, as shown, but many organisms can metabolize the open chain carboxylic acid products of those initial catabolic, or biodegradative, reactions. Intermediary metabolism databases such as KEGG, the Kyoto Encyclopedia of Genes and Genomes, which includes the LIGAND database of enzymes and reactions (Goto et al., 2000), depict this process. For example, one of the metabolic pathways for nitrobenzene is shown on the UM-BBD to yield 2-aminomuconate semialdehyde and the metabolic fate of this latter compound is shown on KEGG (http:// www.genome.ad.jp). It is important to study the pathways of biodegradation to insure that more highly toxic compounds are not generated as the result of microbial metabolism. This concern was raised by the recent observation that the widely used industrial solvent trichloroethylene undergoes reductive dechlorination, generating vinyl chloride as an intermediate (Vogel and McCarty, 1985). See umbbd.ahc.umn.edu/tce2/tce2_map. html for more information about this pathway. Because it is a strong human carcinogen, vinyl chloride is a greater environmental problem than trichloroethylene (Maltoni and Cotti, 1988). So it is not enough to know that a pollutant is disappearing from a given environment; regulatory

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agencies increasingly need to account for its complete environmental fate.

Methodological Advances Relevant to Biodegradation Enrichment Culture In the late 1800s, microbiologists largely focused on the bacteria that cause disease in humans. Isolating the disease-causing bacteria directly from an infected tissue was relatively easy because the infection was largely a monoculture. Thus, plating onto a rich medium might well yield a single organism that could be studied for its disease-causing properties. This contrasted with the situation in a natural soil or water where thousands of different bacteria might well be present in a gram of material. In this case, culturing on a nonselective laboratory medium would yield a complex mixture, difficult to analyze for one particular metabolic trait. Thus, one needed to enrich the mixture to obtain one or a few different types of bacteria. This would simplify the system so that it could be studied productively. As pioneered by the Dutch microbiologist Beijerinck (1901), the enrichment culture technique allowed selective cultivation of one or more bacterial strains obtained from complex environmental mixtures. Assume that one wanted to study the ability of microorganisms in a particular soil to metabolize a given compound. The compound would then be added as the sole carbon, nitrogen or sulfur source to a liquid laboratory medium lacking one of those major elements but containing the others and trace nutrients. The medium would then be inoculated with soil or water, perhaps adding as many as 1011 bacteria. If only a few of the bacteria are able to metabolize the compound to meet their nutritional needs, they will reproduce, or be enriched, selectively. The numbers of these specific bacteria will increase markedly in comparison to what is present in the native soil or water where alternative carbon, nitrogen or sulfur sources are present. With repeated transfer of the enriched microbial mixture into fresh growth medium, the numbers of the preferred bacterium will sometimes increase to the extent that it can be readily isolated. The preferred bacterium is one that can metabolize the given compound, utilize the trace nutrients provided, grow at the temperature used in the laboratory and reproduce quickly. In this context, one may not necessarily obtain the bacterium most prevalent in the original sampled environment. For this reason, some people have

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CHAPTER 1.29

criticized the practice of obtaining and characterizing prokaryotes in pure culture as unsuited to yielding insights into what occurs in nature. But think of the difficulties inherent in trying to learn the details of biodegradation in a complex milieu such as soil. The metabolism of a particular compound might be inferred if it is disappearing from soil, but one has to rule out abiotic reactions in soil and soil can be difficult to sterilize. If the compound is available in a radiolabelled form, an accumulating intermediate may be obtained if it is stable in soil. This may or may not yield insights into metabolic pathways. But one would be hard-pressed to learn about the other metabolites, the enzymes, genes and specific microorganisms involved. In short, without obtaining pure cultures, one could learn whether a compound is metabolized but little about the molecular details. The use of prokaryote pure cultures, many of which have been obtained by enrichment culture, has been instrumental in the identification of the many novel enzymes catalyzing metabolic transformations that drive the carbon, nitrogen and sulfur cycles of Earth. In turn, the corresponding genes have been identified; at first these were identified singly, and now wholesale as the result of genome sequencing efforts, which focused initially on prokaryotes because of their relatively small genome size. Without the development of enrichment culture, we would know far less about the Earth’s biological cycles, the catalytic diversity of the planet and microbial phylogenetic diversity.

per million when coupled with oxygen-scrubbers for the gas mixtures and catalyst cartridges inside the anaerobic chamber. Anaerobic biodegradation is also difficult to study in another context. Anaerobic enrichment cultures may initially show very long lag phases, perhaps six months or one year, before significant biodegradation occurs. Upon repeated transfer, the lag phase often shortens continually. Still, many years may be required to achieve significantly rapid rates of biodegradation and those may never approach the rates of comparable aerobic biodegradation. In most cases, a definitive explanation for the lag phase phenomenon is lacking. It is these kinds of impediments which have skewed the focus of laboratory studies in biodegradation toward the fast-growing aerobic prokaryotes such as Pseudomonas species, which can be grown overnight with simple equipment and typically yield high cell densities. Despite this, anaerobes offer rewards to those who persevere by providing for the discovery of the most novel biochemical reactions on Earth. Some of these reactions have recently been elucidated. For example, bacteria are now known to catabolize toluene anaerobically. They initiate attack on the benzylic carbon via a radical mechanism that generates a new carbon-carbon bond to form benzylsuccinate as the first metabolite (Leuthner et al., 1998). Others, such as the anaerobic formation of methane from long-chain alkanes (Zengler et al., 1999), remain obscure biochemically.

Anaerobic Culturing Methods and Biodegradation

Analytical Chemistry

Most of our early knowledge on biodegradation derived from studies on aerobic or facultative bacteria. This reflected the comparative ease of studying aerobic versus anaerobic bacteria. Anaerobic conditions were fairly easy to maintain with mixed-cultures because facultative organisms would consume oxygen and thus allow strict anaerobes to survive. Obtaining strict anaerobes in pure culture, and elucidating the novel biochemical reactions they catalyze, required the development of specialized techniques (Barker, 1940; Hungate, 1985). Several decades ago, microbiologists used such techniques as roll-tubes to cultivate strictly anaerobic prokaryotes such as methanogenic bacteria. More recently, people routinely began using crimp-sealed, septum-plugged bottles for liquid cultures and putting Petri plates into anaerobic chambers containing an inert gas such as helium or argon. The latter can routinely be maintained at oxygen levels of around one part

Chemical methods for analyzing organic compounds have improved enormously since the late 1800s when use of enrichment culture methods began. Thus, obtaining pure cultures has gone hand in hand with new methods for analyzing the intermediates and products of their metabolism. Thus, one might anticipate a biodegradative metabolic pathway based on chemical logic, obtain authentic chemical standards, and screen for the presence of such compounds in growth cultures of the microbial isolate. But how does one screen for the compound(s)? Chromatography, coupled to the use of authentically synthesized standard compounds, has been a powerful method for studying biodegradation over the last century, and it remains so today. There have been big advances in the science of chromatographic separations. A century ago, thin-layer chromatography (TLC) was state of the art. Later, gas chromatography (GC) provided better resolution and most recently, high pressure liquid chromatography (HPLC) gives

CHAPTER 1.29

excellent resolution and the ability to capture and further analyze compounds. Identification of compounds in complex mixtures is being aided by new developments in mass spectrometry (MS), which may be coupled with isotopic labelling for additional power. Similarly, with high-field nuclear magnetic resonance (NMR) spectroscopy, and the use of specifically labelled 13C-compounds becoming increasingly available, it is now feasible to monitor metabolism in situ (Sauer et al., 1999). This, in turn, may lead to a revolution in environmental microbiology, in which systems more resembling natural systems can be analyzed with respect to biodegradation.

Whole Genome Sequencing and Analysis Biodegradative genes have been identified, usually by transferring the DNA containing a specific gene(s) from a pure culture environmental isolate into Escherichia coli for sequencing and functional expression. With a substantial set of genes available, it is becoming routine to screen soils for the presence of homologous genes that might be involved in identical or similar biodegradative reactions. This gives insights into the environmental prevalence of certain biodegradative genes. In the last several years, DNA sequencing techniques have advanced to the point that we can readily sequence entire prokaryote genomes (Nelson et al., 2000). With appropriate annotation techniques, this can provide insight into the metabolic pathways encoded by the genes. Theoretically, an organism’s entire network of metabolism can be deduced. In practice, deducing metabolism is an imperfect task. Consider that the complete genomic DNA sequence of Escherichia coli, the most intensely studied biochemical entity on Earth, yielded 38% of the coding regions having unknown function (Blattner et al., 1997). If there are gaps in the metabolic map of E. coli, there will be many more as we proceed to sequence the genomic DNA of soil isolates important in biodegradation. Despite this caveat, it is exciting to contemplate the explosive increase in obtaining complete genome sequences for an expanding array of prokaryotes. This will spur a resurgence of interest in comparative biochemistry, with an attendant interest in new “exotic” genes. I predict that a significant number of the newly discovered gene functions in soil Eubacteria will be involved in the biodegradation of organic compounds. This will enhance interest in biodegradation and microbial biocatalysis, in general, as the era of functional genomics comes into full swing.

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The Prokaryotes of Biodegradation Our current perspective on the microorganisms of biodegradation derives largely from enrichment culture methods, isolating pure cultures and studying the individual reactions of biodegradation. Table 1 shows the list of prokaryotes and compounds they degrade; the biodegradative pathways each of them initiate are depicted in the UM-BBD. An analogous microorganism index can be found at umbbd.ahc.umn.edu (UM-BBD). The UM-BBD Microorganism Index has links for the entries containing both genus and species names to the corresponding entries on websites maintained by the American Type Culture Collection (ATCC) or the Deutsche Sammlung von Mikroorganismen und Zellkulturen (DSMZ). Another excellent web resource that deals with microorganisms which are important in biodegradation is the Biodegradative Strain Database (BSD) bsd.cme.msu.edu maintained at Michigan State University by John Urbance, Jim Cole and Jim Tiedje. The BSD microorganism listings, in turn, link to http://www.cme.msu.edu (Ribosomal RNA Database) and to biodegradative pathways maintained on the UM-BBD. Several trends are apparent from perusing the data in Table 1. First, biodegradative capabilities are widespread phylogenically within the Proteobacteria, high G+C Gram-positive bacteria, and Flavobacterium in the Cytophageles-green sulfur bacteria. As discussed in the section above, this reflects the facile transfer of genes, especially those that might be contained on plasmids or flanked by transposable elements. Second, there are several genera of bacteria, which have emerged repeatedly as having diverse catabolism, particularly with starting compounds we think of as metabolically unusual, such as synthetic industrially relevant organic compounds. The latter include herbicides, insecticides, industrial solvents and synthetic intermediates. As illustrated in Table 1, the following genera are particularly well represented: Arthrobacter, Burkholderia, Pseudomonas and Rhodococcus. The caveat to these observations is that we have largely depicted biodegradation pathways catalyzed by prokaryotes, which have been obtained in pure culture via enrichment culture. Thus, we have selectively depicted microorganisms that grow well under conditions typically used for enrichment culture and the maintenance of pure culture isolates in the laboratory. These microbial strains may only reflect some fraction, perhaps a small fraction, of the prokaryotes that actively carry out biodegradation in the soils and waters of the Earth. The complete genome sequencing of both pure culture bacteria

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CHAPTER 1.29

Table 1. Prokaryote genera identified in biodegradation and the compounds they metabolize. a Prokaryote genus Acetobacterium Achromobacter Acinetobacter Actinomycetes Aeromonas Agrobacterium Alcaligenes

Ancylobacter

Arthrobacter

Azoarcus Azotobacter Bacillus Beijerinckia Brevibacterium Brevundimonas Burkholderia

Clavibacter Clostridium Chelatobacter

Compound undergoing biodegradation Triethanolamine Carbon tetrachloride 2,4-Dichlorobenzoate Cyclohexanol 2-Chloro-N-isopropylacetanilide 2,4,6-Trinitrotoluene (TNT) Phenanthrene Glyphosate 1,2,3-Tribromopropane Atrazine 2,4-Dichlorobenzoate 2,4-Dichlorophenoxyacetic acid (2,4-D) 2,4-Dichlorobenzoate 2-Aminobenzenesulfonate Toluene-4-sulfonate Atrazine 1,2-Dichloroethane 2,4-Dichlorophenoxyacetic acid 4-Nitrophenol 1,3-Dichloro-2-propanol Tyrosine 2,4-Dichlorobenzoate Glyphosate Parathion 2,4-Dichlorophenoxyacetic acid (2,4-D) 4-Nitrophenol Octamethylcyclotetrasiloxane Iprodione 1,3-Dichloro-2-propanol Fluorene Tyrosine 2-4-Dichlorobenzoate Glyphosate Methyl tert-butyl ether Nicotine 2-Aminobenzoate Phenanthrene Parathion Benzoate Toluene 2,4-Dichlorophenoxyacetic acid (2,4-D) Thiocyanate 2,4,6-Trinitrotoluene 2-Phenylacetaldoxime Xylene Dibenzofuran Nitrobenzene Parathion 2,4-Dichlorophenoxyacetic acid (2,4-D) 1,2,4-Trichlorobenzene Phthalates Benzoate Pentachlorophenol 2,4,5-Trichlorophenoxyacetic acid (2,4,5-T) 3-Chloroacrylic acid Toluene Trichloroethylene o-Xylene 2,4-Dichlorobenzoate Atrazine 2,4,6-Trirtotoluene (TNT) Phenol Nitrilotriacetate

CHAPTER 1.29

The Metabolic Pathways of Biodegradation

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Table 1. Continued Prokaryote genus Comamonas

Corynebacterium Dehalobacter Dehalococcoides Dehalospirillum Desulfitobacterium Desulfobacterium Desulfovibrio Enterobacter Escherichia Eubacterium Exophiala Fiavobacterium

Hydrogenophaga Hyphomicrobium Klebsiella

Lactobacillus Methanobacterium Methanosarcina Methylobacterium Methylococcus Methylophilus Methylosinus Methylosulfonomonas Moorella Moraxella Mycobacterium Myrothecium Neurospora Nirtosomonas Nocaradia Pelobacter Proteus Pseudomonas

Compound undergoing biodegradation Nirtobenzene 3-Methylquinoline Phthalates Toluene-4-sulfonate 1,3-Dichloro-2-propanol 2,4-Dichlorobenzoate Tetrachloroethene Tetrachloroethene Tetrachloroethene Tetrachloroethene Carbon tetrachloride 2,4,6-Trinitrotoluene (TNT) Glyphosate 1,1,1-Trichloro-2,2-bis-(4-chlorophenyl)ethane (DDT) Pentaerythritol tetranitrate 3-Phenylpropionate Arsonoacetate Phenylmercuric chloride Gallate Styrene Bromoxynil 2,4-Dichlorophenoxyacetic acid (2,4-D) Glyphosate Parathion Pentachlorophenol 4-Carboxy-4¢-sulfoazobenzene Dichloromethane Dimethyl sulfoxide Benzonitrile Bromoxynil Acetylene 1,1,1-Trichloro-2,2-bis-(4¢-chlorophenyl)ethane (DDT) 2,4,6-Trinitrotoluene (TNT) Carbon tetrachloride Tetrachloroethene Carbon tetrachloride Dichloromethane Methyl tert-butyl ether Thiocyanate Dichloromethane Trichloroethylene Methanesulfonic Acid Carbon tetrachloride 4-Nitrophenol 2-Chloro-N-isopropylacetanilide Naphthalenesulfonates Methyl tert-butyl ether Cyanamide 2-Nitropropane Methyl fluoride Dimethyl ether Methyl tert-butyl ether Parathion Methyl ethyl ketone Acetylene 1,1,1-Trichloro-2,2-bis-(4-chlorophenyl)ethane (DDT) Acrylonitrile 2-Aminobenzoate 1,3-Dichloropropene Dichloromethane Dimethyl sulfoxide Carbazole (continued)

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CHAPTER 1.29

Table 1. Continued Prokaryote genus

Ralstorsa Rhodobacter Rhodococcus

Compound undergoing biodegradation Benzoate p-Xylene p-Cymene Carbon tetrachloride Fluorene Adamantanone 3-Chloroacrylic Acid 2-Chloro-N-isopropylacetanilide 1,4-Dichlorobenzene Parathion Nitroglycerin Toluene Octane Nitrobenzene 4-Chlorobiphenyl Dibenzothiophene Orcinol Xylene Ethylbenzene Mandelate Styrene Trichloroethylene Toluene-4-sulfonate m-Xylene Atrazine Naphthalenesulfonates 2,4-Dichlorobenzoate Chlorobenzene 2-Aminobenzoic Acid 4-Chlorobiphenyl Ethylbenzene Naphthalene Chlorobenzene 1-Aminocyclopropane-1-carboxylate Biphenyl Caprolactam Phenanthrene 1,1,1-Trichloro-2,2-bis-(4-chlorophenyl)ethane (DDT) 2,4,6-Trinitrotoluene m-Cresol Thiocyanate Phenylmercuric chloride n-Octane Dodecyl Sulfate Bromoxynil Dibenzothiophene 2,4-Dichlorobenzoate Mandelate Methyl tert-butyl ether (+)-Camphor 2,4-Dichlorophenoxyacetic Acid Atrazine 1,1,1-Trichloro-2,2-bis(4¢-chlorophenyl)ethane (DDT) Dimetyl sulfoxide Acetylene Atrazine Acrylonitrile Methyl tert-butyl ether Cyclohexanol Bromoxynil Styrene Tetrahydrofuran Benzonitrile

CHAPTER 1.29

The Metabolic Pathways of Biodegradation

963

Table 1. Continued Prokaryote genus

Rhodopseudomonas Salmonella Sphingomonas

Sporomusa Staphylococcus

Streptomyces Synechococcus Terrabacter Thauera Thiobacillus Xanthobacter

Compound undergoing biodegradation Dibenzothiophene Benzoate 2,4,6-Trinitrotoluene (TNT) n-Octane Dibenzofuran Carbazole g-1,2,3,4,5,6-Hexachlorocyclohexane Dibenzo-p-dioxin Xylenes Tetrachloroethene Dibenzofuran 2,4,6-Trinitrotoluene Arsonoacetate Fluorene Atrazine Phenanthrene 1,1,1-Trichloso-2,2-bis-(4-chlorophenyl)ethane (DDT) Phenanthrene Dibenzofuran Toluene Benzoate Phenol Thiocyanate 1,2-Dichloroethane 1,4-Dichlorobenzene 2-Chloro-N-isopropylacetanilide 2-Nitropropane Propylene

a

A similar list with links to the metabolic pathways can be obtained on the UM-BBD at http://umbbd.ahc.umn.edu/search/ micro.html

and genomic DNA from soil, the so-called “soil metagenome” (Rondon et al., 2000), may help address this question by helping unveil what percentage of the total genome of a given organism functions in non-intermediary catabolic metabolism. In another example, culture-independent molecular methods were used to analyze microbial communities in an aquifer contaminated with hydrocarbons and chlorinated solvents in which active biodegradation was occurring (Dojka et al., 1998). In that study, 16S rRNA sequences were determined for 21 bacterial members of the consortium, belonging to four recently described divisions of bacteria for which there are no cultivated representatives. Moreover, two particularly abundant 16S rRNA sequence types were implicated in the overall hydrocarbon metabolism. They were members of the genera Syntrophus and Methanosaeta, both of which were proposed to participate in aceticlastic methanogenesis at the end of the catabolic food chain. In parallel with molecular non-culture methods, the well-established methods of enrichment culture are more frequently being applied under anaerobic and other nonstandard conditions in an effort to obtain novel microbial types. This

approach also suggests that biodegradative capabilities are more widespread in the microbial world than has been appreciated by some. For example, halophiles have been identified which metabolize nitroarenes, and members of the Heliobacterium group are known that catabolize polychlorinated biphenyls (PCBs) and chlorophenols (Table 2). These and other recent observations are expanding the taxonomic range of bacteria that catabolize environmental pollutants. Further experiments are likely to expand this further.

Themes in Biodegradation Pathways Occurrence of Similar Pathways in Divergent Prokaryotes Gene transfer amongst prokaryotes is quite facile, and our appreciation of this seems to be increasing all the time. The genes most prone to transfer are those conferring survival advantage only under specialized conditions, the so-called “dispensible genes.” Principal among those

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CHAPTER 1.29

Table 2. Microbes recently identified as organic pollutant biodegraders, but falling outside of the prokaryotic groupings typically isolated for studies on biodegradation. Prokaryotea

Taxonomic group

Haloanaerobium praevalens

Haloanaerobiales

Sporohalobacter marismortui

Haloanaerobiales

Borrelia burgdorferi Borrelia hermsii Bacteroides fragilis Desulfitobacterium dehalogenans Desulfitobacterium hafniense Desulfitobacterium dehalogenans a

Substrate

Reference

Spirochaetales

Nitrobenzene o-Nitrophenol m-Nitrophenol p-Nitrophenol Nitroanilines 2,4-Dinitrophenol 2,4-Dinitroaniline Nitrobenzene o-Nitrophenol m-Nitrophenol p-Nitrophenol Nitroanilines 2,4-Dinitrophenol 2,4-Dinitroaniline Benzamides

Oren et al., 1991

Dettori et al., 1995

Cytophagales Heliobacterium Heliobacterium Heliobacterium

Alkylhydroperoxides Polychlorinated-biphenyls 3-Chloro-4-hydroxy-phenylacetate Chorophenols

Rocha et al., 1999 Wiegel et al., 1999 Christiansen et al., 1998 van de Pas et al., 1999

Oren et al., 1991

These bacteria do not belong to the following groups: Proteobacteria, and high and low G+C Gram-positive bacteria.

genes are ones conferring antibiotic resistance, heavy metal resistance or new catabolic activities. These genes are commonly found on plasmids. Many catabolic plasmids have been shown to have a broad host-range and transfer by conjugation in the absence of helper plasmids. Thus, the genes, and the metabolic functions they encode, can show up in diverse prokaryotes. An example will best serve to illustrate this point. In 1995, a Pseudomonas species, denoted strain ADP, was isolated from an enrichment culture in which the herbicide atrazine was supplied as the sole source of nitrogen (Mandelbaum et al., 1995). Subsequent studies over the ensuing three years elucidated the atrazine catabolic pathway and yielded the DNA sequences of the genes encoding the first three metabolic steps (Fig. 1). During the same period, other laboratories isolated atrazine-catabolizing prokaryotes using different enrichment and isolation condi-

CI N NH

tions (Bouquard et al., 1997; de Souza et al., 1998a; de Souza et al., 1998b; Radosevich et al., 1995; Struthers et al., 1998). The bacteria were subjected to taxonomic determination and found to be members of the following genera, respectively: Rhizobium, Agrobacterium, Ralstonia and Clavibacteria. In our laboratory, DNA from each of the distinct atrazine-catabolizing bacteria was prepared (de Souza et al., 1998b). They were each found to contain genes with more than 99% sequence identity to the atrazine genes from the original Pseudomonas sp. ADP isolate. This occurred despite the fact that the organisms were isolated independently in different regions of Earth, by different groups and under different conditions. These observations are consistent with a facile transfer of the atrazine-catabolic genes amongst soil prokaryotes. In another example, illustrated by perusing the UM-BBD, the same organic compound is metabolized by different genera of bacteria via

OH

OH N

N

Atrazine

N

Atz B NH

NH

N

N

Atz B

N NH

Hydroxyatrazine

HO

OH N

N

NH

N-Isopcopylammelide

Fig. 1. Catabolic pathway for the catabolism of atrazine in Pseudomonas sp. ADP.

N

Atz C HO

N N

OH

Cyanuric Acid

CHAPTER 1.29

The Metabolic Pathways of Biodegradation

different intermediates. Examples of starting compounds are: {atrazine {fluorene {glyphosate {nitrobenzene {4-nitrophenol {parathion {phenanthrene {styrene {toluene {2,4,6-trinitrotoluene

Common Themes in Catabolic Pathways Although some catabolic pathways are widespread, there are some correlations between certain types of metabolism and the prokaryotic group catalyzing those reactions. This can be attributed to the compatibility between the catabolic reactions and the core metabolic pathways that the catabolic intermediates feed into. This is particularly well illustrated for chemical compounds which are composed of a single carbon atom or which are readily metabolized to C-1 fragments. Figure 2 shows the C-1 meta-pathway, which is also depicted in the [{umbbd.ahc.umn.edu/c1/ c1_map.html}{UM-BBD}]. At the core of the map are the oxidative and reductive parts of the C-1 metabolic cycle, which is important on a global scale as discussed previously. Anaerobic Archaea (known as methanogens) catalyze the reductive reactions that transform carbon dioxide to methane. These organisms are important members of certain anaerobic consortia involved in the biodegradation of complex organic matter such as cellulosic wastes. Methanogens occupy the end of the anaerobic food chain in the overall biodegradative process. A class of prokaryotes called “methanotrophs” (Fig. 2) carry out the oxidative reactions leading from methane, the most reduced C-1 compound, to carbon dioxide, the most oxidized. Some C-1 oxidizing organisms (known as “methylotrophs”) cannot oxidize methane to methanol, but can carry out the next three oxidative reactions. Methane is a common natural product; it is the main constituent of natural gas. Moreover, data suggests that a majority of the methane generated in lake sediments is oxidized in higher, aerobic levels of the lake by methanotrophs, and thus methane never enters the atmosphere. Methanotrophic and methylotrophic metabolism may be expanded to include a set of oxidative, hydrolytic or thiolytic reactions whereby simple organic structures can be transformed to the methanotrophic intermediates methanol, formaldehyde or formate (Fig. 2). Some of these

965

O CH3P(OH)2

CH3HgCl

CH4 CH3OCH3

CH3OSO3H CH3OH CH3Br

CH3SO3H

CH3NH2 CH2Cl2 HC

O HCHO

H3CSCH3

N HCO2H CO CS2

CO2

HSCN

Fig. 2. Catabolism of organic compounds containing one carbon atom that funnel into the central intermediates of methanotrophy: methane, methanol, formaldehyde and formate.

compounds are natural products; for example dimethyl sulfide, methylamine and methyl chloride. Others are predominantly the products of organic synthesis: dichloromethane, dimethylether and methyl fluoride. Regardless of their origin, these compounds are readily transformable to methanotrophic metabolic intermediates and thus some methanotrophs will grow on them as their sole source of carbon and energy. This catabolic metabolism is not universal, however. Only some small subset of the total set of methanotrophs and methylotrophs will grow on a given compound shown at the periphery of Fig. 2. But methanotrophs and methylotrophs are common in nature and thus dichloromethane, dimethylether and methyl fluoride are generally thought of as being fairly biodegradable. Another common theme is seen in the transformation of the commercially important BTEX compounds (i.e., benzene, toluene, ethylbenzene and xylenes). They are clustered because of their co-occurrence in environmental contamination stemming from spillage of petroleum materials. Because BTEX compounds are structurally analogous to each other, there are commonalities in their metabolism by prokaryotes. Anaerobic metabolism of BTEX compounds has been studied only more recently, and the biochemical basis of the biodegradation reactions is now being revealed. The aerobic metabolism of BTEX compounds is much better studied. For example, see Fig. 3 and (umbbd.ahc.umn.edu/BTEX/

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CHAPTER 1.29 R

OH

OH Catechol

R H R = CH3

Benzene

H OH

Toluene

or

CH3CH2 Ethylbenzene X = CH3

OH

OH

Prokaryota OH

Xylene

OH H cis -Dihydrodiol

Phenol

x

Eukaryota

H

CH3 Fig. 3. Convergence of aerobic pathways for BTEX compounds leads to a catechol intermediate.

BTEX_map.html#aerobic). Almost invariably, oxygenase enzymes initiate the metabolism to produce ring cis-dihydrodiols, phenols, benzyl alcohols and ultimately catechols, which undergo ring cleavage. These alcohol products are all more activated than their aromatic hydrocarbon starting compounds. There are multiple pathways possible but all of them produce catechol intermediates. To follow all known aerobic prokaryotic metabolic pathways for each of the BTEX compounds, follow the links: {Benzene {Toluene {Ethylbenzene {o-Xylene {m-Xylene {p-Xylene The metabolic strategy for BTEX compounds used by aerobic prokaryotes differs from that used by aerobic eukaryotic organisms such as fungi (Cerniglia et al., 1978) and mammals (Jerina et al., 1968). The latter group also uses oxygenase enzymes to attack resonancestabilized aromatic hydrocarbons. However, the initial products of the oxygenase-catalyzed reactions are often arene oxides (Fig. 4). Aromatic alcohols are detected but are shown to largely arise from spontaneous isomerization of the arene oxides and are not direct enzyme products. This contrasts with the prokaryote aromatic ring monooxygenation reactions in which the phenol product is detected directly. For example, with

O H Arene oxide Fig. 4. Divergence in the catabolism of aromatic compounds by Prokaryota and Eukaryota.

toluene catabolism by Burkholderia cepacia G4, the initial reaction product is 2-hydroxytoluene, or o-cresol, exclusively. There was no evidence for the intermediate formation of toluene 2,3-epoxide, which would have isomerized to a mixture of o-cresol and m-cresol. The data do not rule out that an epoxide is an enzyme-bound intermediate that undergoes a controlled isomerization on the enzyme surface. This would be advantageous to the organism, as epoxides are reactive electrophiles and can alkylate proteins and other molecules in the cell. So a high-flux metabolic pathway that produces such a reactive species might well be selected against during evolution. In contrast, the mammalian metabolism of BTEX compounds is largely low flux metabolism, to scavenge stray hydrocarbons that may enter the body. Other enzymes further metabolize the epoxide products to make intermediates that are excreted from the animal. A nonspecific detoxification metabolism such as this may work best when it proceeds through an initial arene oxide intermediate. Acknowledgments. I thank the many talented colleagues with whom I have collaborated on metabolic pathways of biodegradation. Special thanks go to Professor Lynda Ellis for her meticulous work on the University of Minnesota Bio-

CHAPTER 1.29

catalysis/Biodegradation Database, making it a resource that has been worth accessing millions of times by researchers around the world.

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egy for accessing the genetic and functional diversity of uncultured microorganisms. Appl. Environ. Microbiol. 66:2541–2547. Sauer, U., D. R. Lasko, J. Fiaux, M. Hochuli, R. Glaser, T. Szyperski, K. Wuthrich, and J. E. Bailey. 1999. Metabolic flux ratio analysis of genetic and environmental modulations of Escherichia coli central carbon metabolism. J. Bacteriol. 181:6679–6688. Spain, J. C. 1995. In: J. C. Spain (Ed) Biodegradation of Nitroaromatic Compounds. Plenum Press. New York, NY. Strong, L. C., H. McTavish, M. J. Sadowsky, and L. P. Wackett. 2000. Field-scale remediation of atrazinecontaminated soil using recombinantEscherichia coli expressing atrazine chlorohydrolase. Environ. Microbiol. 2:91–98. Struthers, J. K., K. Jayachandran, and T. B. Moorman. 1998. Biodegradation of atrazine by Agrobacterium radiobacter J14a and use of this strain in bioremediation of contaminated soil. Appl. Environ. Microbiol. 64:3368– 3375. Van de Pas, B. A., H. Smidt, W. R. Hagen, J. van der Oost, G. Schraa, A. J. Stams, and W. de Vos. 1999. Purification

CHAPTER 1.29 and molecular characterization of ortho-chlorophenol reductive dehalogenase, a key enzyme of halorespiration in Desulfitobacterium dehalogenans. J. Biol. Chem. 274:20287–20292. Vogel, T. M., and P. L. McCarty. 1985. Biotransformation of tetrachloroethylene to trichloroethylene, dichloroethylene, vinyl chloride, and carbon dioxide under methanogenic conditions. Appl. Environ. Microbiol. 49:1080– 1083. Wagner-Dobler, I., H. von Canstein, Y. Li, K. N. Timmis, and W.-D. Deckwer. 2000. Removal of mercury from chemical wastewater by microorganisms in technical scale. Environ. Sci. Technol. 34:4628–4634. Wiegel, J., X. Zhang, and Q. Wu. 1999. Anaerobic dehalogenation of hydroxylated polychlorinated biphenyls by Desulfitobacterium dehalogenans. Appl. Environ. Microbiol. 65:2217–2221. Winogradsky, S. 1890. Sur les organismes de la nitrification. Compt. Rend. Acad. Sci. 110:1013–1016. Zengler, K., H. H. Richnow, R. Rossello-Mora, W. Michaelis, and F. Widdel. 1999. Methane formation from long-chain alkanes by anaerobic microorganisms. Nature 401:266– 269.

Prokaryotes (2006) 2:969–984 DOI: 10.1007/0-387-30742-7_30

CHAPTER 1.30 c i l i hp i l ak l ao l aH

gn i z i d i xO- ruf l uS

a i re t caB

Haloalkaliphilic Sulfur-Oxidizing Bacteria DIMITRY YU. SOROKIN, HORIA BANCIU, LESLEY A. ROBERTSON AND J. GIJS KUENEN

Introduction

Soda Lakes as a Unique Habitat

Chemolithoautotrophic sulfur-oxidizing bacteria (SOB) play an important role in the element cycling in natural and man-made environments because of their extremely high capacity to transform various sulfur compounds and their contribution to secondary production of organic matter. They are widely distributed in various habitats, associating primarily with sulfideoxygen interface layers, where they successfully compete with chemical sulfide oxidation by oxygen. Energetically, the reaction of complete oxidation of sulfide or thiosulfate to sulfate (8 electrons) is among the most attractive for chemosynthesis, and not surprisingly, sulfur oxidizers can be found in many different groups of prokaryotes. Currently, lithoautotrophic sulfur bacteria are mostly found among the proteobacteria (alpha, beta, gamma and epsilon subdivision; The Colorless Sulfur Bacteria in the second edition; Kuenen and Robertson, 1992; Kelly and Wood, 2000). The currently known exceptions outside the proteobacteria are among the Grampositive bacteria (Sulfobacillus), Crenarchae (Sulfolobus), and deep lineages (Aquificalis; The Chemolithotrophic Prokaryotes in this Volume). According to their response to pH, the known sulfur-oxidizing species include acidophiles (optimum pH 1.5 M total Na+. Therefore the most important selective force favoring survival of alkaliphilic sulfur bacteria in soda lakes appeared to be the salt concentration (Table 3).

Ecophysiology of Aerobic Haloalkaliphilic SulfurOxidizing Bacteria The main properties of the two groups of haloalkaliphilic sulfur bacteria are presented in Tables 4 and 5. The Thioalkalimicrobium group is represented by highly specialized, low-salt tolerant, fast-growing and low-yield strains with extremely high sulfide- and thiosulfate-oxidizing activity. In contrast, the Thioalkalivibrio group is more physiologically diverse and accommodates slowly growing organisms with more efficient substrate conversion. These are, in general, more salt-tolerant, with many strains able to grow in saturated soda brines. This group uniformly synthesized a membrane-bound yellow pigment not found in the low-salt tolerant Thioalkalivibrio strains. This pigment is a 23-carbon polyene compound with a structure unlike that of any known bacterial pigments (Takaichi et al., 2004). Although its complete formula and function is not yet completely understood, it is clearly

CHAPTER 1.30

Haloalkaliphilic Sulfur-Oxidizing Bacteria

973

0.05

98

Tv  ALMg 2 Uncultured clone ML623J-18 Tv . AL 2 Tv ALJ 12 Tv ARH 2  T

98

ALJ22 Tv HL17 ALJ 24 AKLD 2 100 Tv ALEN2 Uncultured clone ML635J-54 Tv ARh 1 Tv ALJD 100 Tv ARhD 1 99 A12  DSM 4180 E E 100 100 DSM 2111   DSM 243 DSM 237 E  100 DSM 241 E 9902 100 ATCC 35916 100 DSM 2110  E DSM 244 100 99 HA-1 ! E 96 ATCC 25380 E  100 DSM 12769 EE  2  E MLHE-1 100 34Alc E 100 AHN 1 ALEN 1 E 100 symbiont of Solemya reidi symbiont of Solemya reidi gill 100

100

96

E

DSM 2157 CIP 103589

E

100

97

E 

100

E

E E

100

E

100 100 100

E E E

100



E

E



E

E



E



JB-A1 JCM 7688 ATCC 35932 JB-A2 Ch-1 DSM 5322 DSM 1534 E AL 3 E ALM 1 E AL 7

4210 Fig. 2. Phylogenetic tree demonstrating position of the three new genera of haloalkaliphilic sulfur oxidizing bacteria isolated from the soda lakes. Numbers on the branches indicates bootstrap values (only the highest values are included). Unaffiliated strains among the genus Thioalkalivibrio: extremely salt tolerant strains from Mongolia (ALMg 2) and Kenya (ALJ 15, ALJ 22, and ALJ 24); AKLD 2 is a facultatively anaerobic nitrate-reducing strain from Kulunda. Bar, 5% sequence divergence.

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CHAPTER 1.30

Table 3. Results of enrichment and isolation of two different types of haloalkaliphilic SOB from soda lakes (pH 10). Low-salt medium (0.6M Na+) Location Lake Hadyn (Tuva) Kunkur Steppe (Siberia) Northeast Mongolia Lake Borzinskoe (Siberia) Kulunda Steppe (Siberia) Kenya (Rift Valley) Egypt (Wadi Natrun) Mono Lake (California)

High-salt medium (2–4M Na+)

MPN

Tm

Tv

MPN

Tm

Tv

nd 106 106 106 108 106 106 nd

1 14 20 0 3 3 4 1

1 4 0 2 6 20 5 0

nd nd 105 107 108 106 106 nd

nd nd 0 0 0 0 0 0

nd nd 20 1 7 5 23 1

Abbreviations: MPN, maximum cell number/cm3 of sediment; Tm, number of isolated Thioalkalimicrobium strains; Tv, number of isolated Thioalkalivibrio strains; and nd, not determined.

Table 4. Properties of haloalkaliphilic SOB from soda lakes. Property DNA G + C mol% Cell morphology Intracellular sulfur globules Carboxysomes Colony morphology pH limits (optimum) Upper temperature limit (∞C) Upper salt limit (M total Na+) Max.specific growth rate (pH 10) Max.growth yield (g of protein/ mol of thiosulfate) Survival during starvation Rates of thiosulfate and sulfide oxidation Rates of sulfur oxidation Sulfur intermediates Denitrification Growth with thiocyanate RuBisCo activity Sulfite-dehydrogenase Dominating cytochromes Cytochrome oxidases N-sources for growth NH3 NO2-, NO3SCNDominant ubiquinone Compatible solutesa Dominant fatty acids in membrane lipidsb Membrane-bound yellow pigment

Thioalkalimicrobium (43 strains)

Thioalkalivibrio (43 strains) 61–66 Mostly vibrios or short spirilla with a single polar flagellum; some strains are nonmotile, barrel-shaped or coccoid +/+/Compact, often with sulfur, often yellowish

47.3–51.2 From rods to spirilla with 1–3 polar flagella

7.5–10.65 (10–10.2) 50 4.3 0.20h-1 6.5

+ Compact or spreading, pink, without sulfur 7.5–10.6 (10) 39 1.5 0.33h-1 3.5

Long Low-moderate

Short Extremely high

Moderate Polysulfide, sulfur +/+/+, type I + c and b o, cbb3, aa3

Very low Sulfite +, type I c cbb3

+ +/+/Q-8 Glycine betaine C16:0, C18:1, and C19-cyclopropyl

+ + Q-8 Ectoine C16:1, C18:1, and C16:0

+/-

-

Symbols and abbreviation: +, present; - absent; +/-, present in some strains; and RuBisCo, ribulose-1,5-bisphosphate carbosylase oxygenase. a Data of E. Galinski. b Data of J. Sinninghe Damste.

CHAPTER 1.30

Haloalkaliphilic Sulfur-Oxidizing Bacteria

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Table 5. Respiratory activity in haloalkaliphilic SOB grown with thiosulfate or thiocyanate at pH 10. Thioalkalimicrobium Substrate

V -2

Thiosulfate (S2O3 ) Sulfide (HS-) Polysulfide (S8-2) Elemental sulfur (S8) Sulfite (SO3-2) Trithionate (S3O6-2) Tetrathionate (S4O6-2) Pentathionate (S5O6-2) Thiocyanate (SCN-) CS2 (carbon disulfide)

Thioalkalivibrio

pH opt

2.5–5.2 9–10 2.3–5.2 9–10 1.1–3.0 10 0–0.2 10 0 n.d. 0 n.d. 0–1.1 9 0 n.d. No growth and respiration

N

V

pH opt

N

40 40 38 40 28 9 40 9

0.15–1.1 0.15–1.5 0.2–0.9 0.08–0.6 0–0.2 0–0.2 0.05–0.5 0.1–0.8 0.09–0.4 0.09

9–10 9–10 10 10–10.5 10 9 9 9 10 10

60 60 55 60 40 20 60 20 9 1

Abbreviations: V, respiration rate, mmol of O2 (mg of protein min)-1; N, number of tested strains; and n.d., not determined.

N Cs a

10 mm S b

b

a

d

c e

c

10 mm

d

f

Pg f

e Fig. 3. Cell morphology of the genus Thioalkalimicrobium. (a, c–e) — total electron microphotographs; (b, f) — thin sections; (a–b) — Tm.aerophilum AL 3; (c) — str.ALJ 14 (Kenya); (d) — Tm.sibiricum AL 7; (e–f) — Tm.cyclicum ALM 2; Cs-carboxysomes; N-nucleoide. Bars: 0.5 mm.

essential for the functioning of these bacteria at extremely high salt and high pH conditions. The typical cell morphology of these bacteria is shown in Figs. 3 and 4. Soda lake sulfur oxidizers differ from all other known sulfur bacteria because of their ability to grow and oxidize sulfur compounds at pH >9. The sodium carbonate-bicarbonate buffer appears to be the most appropriate mineral environment for such bacteria, providing both stable alkalinity and a source of carbon. The buffering capacity of the carbonate system is maximal at

1 mm

N

N g

Cs

h

Fig. 4. Cell morphology of the genus Thioalkalivibrio. (a–d) — phase contrast of extremely salt tolerant isolates from Kulunda, Kenya and Egypt. (e–h) — thin sections of str.AL2, ALJ 15, ALJ 3, ALE 11. Bar (e,g,h) = 0.5 mm; Cs-carboxysome, N-nucleoide, Pg-polyglucose, S-intracellular sulfur.

pH 9.5–10.2. This pH range was suitable for batch cultivation. However, to explore a realistic pattern of the pH-dependence of growth, continuous cultivation under pH-controlled conditions was necessary (Sorokin et al., 2003b). This investigation confirmed the obligately alkaliphilic

CHAPTER 1.30

0.3

3.5

Y

0.25 µ (h–1)

4

µ

3

0.2 0.15

2.5

0.1 2

0.05

Y (g prot/Mole S2O32–)

Thioalkalimicrobium aerophilum AL 3 0.35

7

8

9

10

Tm

3 2 1

Tv

0 7.5

8.5

11

9.5

10.5

11.5

pH

pH

a

Fig. 6. pH profile of the activity of thiosulfate-dependent respiration for Thioalkalimicrobium (Tm) and Thioalkalivibrio (Tv).

Thioalkalivibrio versutus AL 2 6

µ

Y

5

0.2

4

0.15

3 0.1

2

0.05

1

0

0 7

8

9

10

Y (g prot/Mole S2O32–)

0.25

m (h–1)

4

6.5

1.5

0

b

Activity µ mol/(mg prot min)

D.Y. Sorokin et al.

11

pH

Fig. 5. pH profiles for growth rate (m) and growth yield (Y) of alkaliphilic sulfur oxidizing bacteria measured in pHcontrolled thiosulfate-limited continuous culture (0.6 M total Na+).

Growth rate, % of maximum

976

120 100 80 Tv3

60 40

Tv2

20 Tm, Tv1

0 0

0.5

1

nature of the representative strains of Thioalkalimicrobium and Thioalkalivibrio and, for the first time, demonstrated that chemolithoautotrophic bacteria are capable of stable growth at pH >10 (Fig. 5). Both the growth rate and the growth yield of the soda lake isolates were maximum at pH values around 10. The maximum pH for growth registered in chemostat cultures was 10.6. On the other hand, the pH for maximum respiratory activity was at least 11.0 (Fig. 6). The failure to grow at pH >10.6 might be explained by an anabolic constraint, most probably the unavailability of carbon in the form of CO3–2, as has been suggested previously for alkaliphilic cyanobacteria (Kaplan et al., 1982). Another important environmental factor in the selection of a particular type of SOB is the total salt content. Although all strains isolated from the soda lakes belonged to the haloalkaliphiles, three different subgroups can be identified on the basis of their salt tolerance and requirement (Fig. 7a). 1) All Thioalkalimicrobium and some of the Thioalkalivibrio isolates

Growth rate, % of maximum

a

2

Total

Na+,

2.5

3

3.5

4

M

120 100 80 ALE 20

60 40

ALE 10

20 ALJ 15

0 0

b

1.5

0.5

1

1.5

2

2.5

3

ALE 28

3.5

4

Cl–, M

Fig. 7. Influence of sodium carbonate concentration (a) and Cl- (b) on growth of different subgroups of extremely salttolerant alkaliphilic sulfur oxidizing bacteria at pH 10. The background Cl- concentration in (a) was 0.1 M, the background carbonate concentration in (b) was 0.5 M of total Na+. Tm, Thioalkalimicrobium; Tv1, low-salt tolerant Thioalkalivibrio; Tv2, extremely natronotolerant Thioalkalivibrio; Tv3, extremely natronophilic Thioalkalivibrio; ALJ 15, Kenyan natronophilic Thioalkalivibrio strain; and ALE 10, ALE 20 and ALE 28, haloalkaliphilic Thioalkalivibrio isolates from Wadi Natrun in Egypt.

CHAPTER 1.30

belong to a moderately salt tolerant type, being able to grow in up to 1.2–1.5 M total Na+. They originated mostly from the hyposaline lakes and were isolated on medium containing a low salt concentration. 2) The biggest group of the Thioalkalivibrio isolates was extremely salt-tolerant and able to grow in saturated soda brines (4– 4.5 M Na+). However, most of them grew optimally at moderate salt concentrations (0.5–1 M Na+). 3) The third type consisting of only a few isolates was the true extreme halophiles that cannot grow at salt concentrations below 1 M Na+. All extremely salt tolerant Thioalkalivibrio strains were isolated from hypersaline soda lakes, mostly in Mongolia (Sorokin et al., 2004a) and Egypt (our unpublished results). The extreme halophiles were also the most thermotolerant, some being able to grow up to 50∞C. Continuous culture experiments with one of these isolates, Thioalkalivibrio versutus ALJ 15, demonstrated its excellent adaptation to doubly extreme conditions (Banciu et al., 2004b). Its growth rate and growth yield in soda brine at pH 10 and 4 M Na+ were only 3 and 2 times lower, respectively, than found at 0.6 M Na+. Not only the total sodium concentration was important for optimal growth and activity, but also the anionic composition of the sodium salts. In particular, the ratio of carbonates to Cl– was critical. Most of the extremely salt tolerant Thioalkalivibrio (isolates from Kenyan, Mongolian and Kulunda soda lakes) were able to grow in pure soda brines without Cl–, but for maximum growth they required 0.1–0.5 M Cl–. Higher concentrations of Cl– inhibited growth, resulting in complete inhibition at >2 M Cl–. In contrast, the strains isolated from the NaCldominated Wadi Natrun lakes, had an obligate requirement for 0.5 M Cl–, grew optimally at 1– 2 M Cl– and still grew at 3–3.5 M NaCl in the presence of only 0.5 M Na+ and carbonate to maintain an alkaline pH and provide the carbon source (Fig. 7b). The latter strains can be regarded as true haloalkaliphiles, while the dominant subgroup of the extremely salt-tolerant Thioalkalivibrio strains does not fit this term. We suggest calling such bacteria “natronophiles”—the soda-loving bacteria. Under low-salt conditions, both Thioalkalimicrobium and Thioalkalivibrio representatives developed in some of the enrichment cultures. Competition experiments in thiosulfate-limited continuous culture at low salt and high pH conditions demonstrated that Thioalkalivibrio has a competitive advantage over Thioalkalimicrobium at extremely low dilution rates (60%). On the other hand,DNA homology between various “geographic species” was usually low (15–50%) despite their very similar phenotypes. At this moment, several molecular fingerprinting techniques are being employed to solve this problem. Molecular techniques are now being used to improve detection of haloalkaliphilic sulfur bacteria in mixed populations in soda lake sediments and sulfide-removing bioreactors. So far, successful oligonucleotide probes have been designed forfluorescence in situ hybridization and polymerase chain reaction detection of the representatives of the Thioalkalivibrio (G. Muyzer and D. Sorokin, unpublished results). Acknowledgments. We would like to thank our colleagues B.E. Jones and G.A. Zavarzin for making possible our work with their samples from the Kenyan soda lakes. We are also grateful to T.P. Tourova and A.M. Lysenko for help in genetic work, to M. Mityushina and K. Sjollema for electron microscopy and to E. Galinski and J. Sinninghe Damste for analyses of compatible solutes and fatty acids.

Literature Cited Antipov, A. N., D. Y. Sorokin, N. P. L’vov, and J. G. Kuenen. 2003. New enzyme belonging to the family of molybdenum-free nitrate reductases. Biochem. J. 369:185–189. Banciu, H., D. Y. Sorokin, E. A. Galinski, G. Muyzer, R. Kleerebezem, and J. G. Kuenen. 2004a. Thioalkalivibrio halophilus sp. nov., a novel obligately chemolithoautotrophic facultatively alkaliphilic and extremely salttolerant sulfur-oxidizing bacterium from a hypersaline alkaline lake. Extremophiles 8:325–334. Banciu, H., D. Y. Sorokin, R. Kleerebezem, G. Muyzer, E. A. Galinski, and J. G. Kuenen. 2004b. Influence of sodium on the growth of haloalkaliphilic sulfur-oxidizing bacterium Thioalkalivibrio versutus strain ALJ 15 in continuous culture. Extremophiles 8:185–192. Baumgarte, S. 2003. Microbial Diversity of Soda Lake Habitats [PhD thesis]. Carolo-Wilhelmina University. Braunschweig, Germany. 79–81. De Kruyff, C. D., J. I. van der Walt, and H. M. Schwartz. 1957. The utilization of thiocyanate and nitrate by thiobacilli. Ant. v. Leeuwenhoek 23:305–316. Eugster, H. P. 1970. Chemistry and origins of the brines of Lake Magadi. Mineral. Soc. Am., Spec. Publ. 3:215–235. Friedrich, C. G., D. Rother, F. Bardischewsky, A. Quentmeier, and J. Fischer. 2001. Oxidation of reduced inorganic sulfur compounds by bacteria: Emergence of

CHAPTER 1.30 a common mechanism?. Appl. Environ. Microbiol. 67:2873–2882. Gorlenko, V. M., B. B. Namsaraev, A. V. Kulyrova, D. G. Zavarzina, and T. N. Zhilina. 1999. Activity of sulfatereducing bacteria in the sediments of the soda lakes in south-east Transbaikal area. Microbiology 68:580–586. Grant, W. D., and B. J. Tindall. 1986. The alkaline saline environment. In: R. A. Herbert and G. A. Codd (Eds.) Microbes in Extreme Environments. Academic Press. London, UK. 25–54. Horikoshi, K. 1991. Microorganisms in Alkaline Environments. Kodansha. Tokyo, Japan. Humayoun, S. B., N. Bano, and J. T. Hollibaugh. 2003. Depth distribution of microbial diversity in Mono Lake, a meromictic soda lake in California. Appl. Environ. Microbiol. 69:1030–1042. Imhoff, J. F., H. G. Sahl, G. S. H. Soliman, and H. G. Trüper. 1979. The Wadi Natrun: Chemical composition and microbial mass developments in alkaline brines of eutrophic desert lakes. Geomicrobiol. J. 1:219–234. Imhoff, J. F., and J. Süling. 1996. The phylogenetic relationship among Ectothiorhodospiraceae: a reevaluation of their taxonomy on the basis of 16S rDNA analyses. Arch. Microbiol. 165:106–113. Isachenko, B. L. 1951. Chloride, sulfate and soda lakes of Kulunda steppe and its biogenic processes [in Russian]. Selected Works. Academy of Sciences USSR. Leningrad, Russia. 2:143–162. Jones, B. F., H. P. Eugster, and S. L. Rettig. 1977. Hydrochemistry of the Lake Magadi basin, Kenya. Geochim. Cosmochim. Acta 41:53–72. Jones, B. E., W. D. Grant, A. W. Duckworth, and G. G. Owenson. 1998. Microbial diversity of soda lakes. Extremophiles 2:191–200. Kaplan, A., D. Zenvirth, L. Reinhold, and J. A. Berry. 1982. Involvement of a primary electrogenic pump in the mechanism of HCO3- uptake by the cyanobacterium Anabaena variabilis. Plant Physiol. 69:978–982. Kappler, U., and C. Dahl. 2001. Enzymology and molecular biology of sulfite oxidation. FEMS Microbiol. Lett. 203:1–9. Kelly, D. P., J. K. Shergill, W.-P. Lu, and A. P. Wood. 1997. Oxidative metabolism of inorganic sulfur compounds by bacteria. Ant. v. Leeuwenhoek 71:95–107. 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. Kuenen, J. G., L. A. Robertson, and O. H. Tuovinen. 1992. The genera Thiobacillus, Thiomicrospira and Thiosphaera. In: A. Balows, H. G. Trüper, M. Dworkin, W. Harder, and K.-H. Schleifer (Eds.) The Prokaryotes. Springer. New York, NY. 3:2638–2657. Loiko, N. G., V. S. Soina, D. Y. Sorokin, L. L. Mityushina, and G. I. El’-Registan. 2003. Production of resting forms by the Gram-negative chemolithoautotrophic bacteria Thioalkalivibrio versutus and Thioalkalimicrobium aerophilum. Microbiology 72:285–294. Nelson, D. C., B. B. Jorgensen, and N. P. Revsbech. 1986. Growth pattern and yield of a chemoautotrophic Beggiatoa sp. in oxygen-sulfide microgradients. Appl. Environ. Microbiol. 52:225–233. Pfennig, N., and K. D. Lippert. 1966. Über das Vitamin B12bedürfnis phototropher Schwefelbakterien. Arch. Mikrobiol 55:245–256.

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Pronk, J. T., R. Meulenburg, W. Hazeu, P. Bos, and J. G. Kuenen. 1990. Oxidation of reduced inorganic sulfur compounds by acidophilic thiobacilli. FEMS Microbiol. Rev. 75:293–306. Rees, H. C., W. D. Grant, B. E. Jones, and S. Heaphy. 2004. Diversity of Kenyan soda lake alkaliphiles assessed by molecular methods. (Online). Extremophiles 8. Robertson, L. A., and J. G. Kuenen. 1992. The use of natural bacterial populations for the treatment of sulfurcontaining wastewater. Biodegradation 3:239–254. Sorokin, D. Y., A. de Jong, L. A. Robertson, and J. G. Kuenen. 1998. Purification and partial characterizaton of sulfide dehydrogenase from alkaliphilic obligately autotrophic sulfur oxidizing bacterium. FEBS Lett. 427:11–14. Sorokin, D. Y., L. A. Robertson, and J. G. Kuenen. 2000. Isolation and characterization of obligately chemolithoautotrophic alkaliphilic sulfur-oxidizing bacteria. Ant. v. Leeuwenhoek 77:251–260. Sorokin D. Y., J. G. Kuenen, and M. Jetten. 2001a. Denitrification at extremely alkaline conditions in obligately autotrophic alkaliphilic sulfur-oxidizing bacterium Hioalkalivibrio denitrificans. Arch. Microbiol. 175:94– 101. Sorokin D. Y., T. P. Tourova, A. M. Lysenko, and J. G. Kuenen. 2001b. Microbial thiocyanate utilization under highly alkaline conditions. Appl. Environ. Microbiol. 67:528–538. Sorokin D. Y., A. M. Lysenko, L. L. Mityushina, T. P. Tourova, B. E. Jones, F. A. Rainey, L. A. Robertson, and J. G. Kuenen. 2001c. 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. Sorokin, D. Y., V. M. Gorlenko, T. P. Tourova, T. V. Kolganova, A. I. Tsapin, K. H. Nealson, and J. G. Kuenen. 2002a. Thioalkalimicrobium cyclicum sp. nov. and Thioalkalivibrio jannaschii sp. nov., new species of alkaliphilic, obligately chemolithoautotrophic sulfuroxidizing bacteria from a hypersaline alkaline Mono Lake (California). Int. J. Syst. Evol. Microbiol. 52:913– 920. Sorokin D. Y., T. P. Tourova, T. V. Kolganova, K. A. Sjollema, and J. G. Kuenen. 2002b. Thioalkalispira microaerophila gen. nov., sp. nov., a novel lithoautotrophic, sulfuroxidizing bacterium from a soda lake. Int. J. Syst. Evol. Microbiol. 52:2175–2182. Sorokin, D. Y., T. P. Tourova, A. M. Lysenko, L. L. Mityushina, and J. G. Kuenen. 2002c. Thioalkalivibrio thiocyanooxidans sp. nov. and Thioalkalivibrio paradoxus sp. nov., novel alkaliphilic, obligately autotrophic, sulfur-oxidizing bacteria from the soda lakes able to grow with thiocyanate. Int. J. Syst. Evol. Microbiol. 52:657– 664. Sorokin, D. Y., A. N. Antipov, and J. G. Kuenen. 2003a. Complete denitrification in coculture of obligately chemolithoautotrophic haloalkaliphilic sulfur-oxidizing bacteria from a hypersaline soda lake. Arch. Microbiol. 180:127–133. Sorokin, D. Y., H. Banciu, M. van Loosdrecht, and J. G. Kuenen. 2003b. Growth physiology and competitive interaction of obligately chemolithoautotrophic, haloal-

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kaliphilic, sulfur-oxidizing bacteria from soda lakes. Extremophiles 7:195–203. Sorokin, D. Y., T. P. Tourova, K. A. Sjollema, and J. G. Kuenen. 2003c. Thioalkalivibrio nitratireducens sp. nov., a nitrate-reducing member of an autotrophic denitrifying consortium from a soda lake. Int. J. Syst. Evol. Microbiol. 53:1779–1783. Sorokin, D. Y., V. M. Gorlenko, B. B. Namsaraev, Z. B. Namsaraev, A. M. Lysenko, B. T. Eshinimaev, V. N. Khmelenina, Y. A. Trotsenko, and J. G. Kuenen. 2004a. Prokaryotic communities of the north-eastern Mongolian soda lakes. Hydrobiologia 522:235–248. Sorokin, D. Y., T. P. Tourova, A. N. Antipov, G. Muyzer, and J. G. Kuenen. 2004b. Anaerobic growth of the haloalkaliphilic denitrifying sulphur-oxidising bacterium Thialkalivibrio thiocyanodenitrificans sp. nov. with thiocyanate. Microbiology (UK) 150:2435–2442. Takaichi, S., T. Maoka, N. Akimoto, D. Y. Sorokin, H. Banciu, and J. G. Kuenen. 2004. Two novel yellow pigments natronochrome and chloronatronochrome from the natrono(alkali)philic sulfur-oxidizing bacterium Thialkalivibrio versutus ALJ 15. Tetrahedron Lett. 45(45):4303–4305.

CHAPTER 1.30 Tindall, B. J. 1988. Procaryotic life in the alkaline, saline, athalassic environment. In: F. Rodriguez-Valera (Ed.) Halophilic Bacteria. CRC Press. Boca Raton, FL. 31– 67. Visser, J. M., G. A. H. de Jong, L. A. Robertson, and J. G. Kuenen. 1997. A novel membrane-bound flavocytochrome c sulfide dehydrogenase from the colorless sulfur bacterium Thiobacillus sp. W5. Arch. Microbiol. 167:295–301. Youatt, J. B. 1954. Studies on the metabolism of Thiobacillus thiocyanooxidans. J. Gen. Microbiol. 11:139–149. Zavarzin, G. A., T. N. Zhilina, and V. V. Kevbrin. 1999. The alkaliphilic microbial community and its functional diversity. Mikrobiology 68:503–521. Zavarzin, G. A., and T. N. Zhilina. 2000. Anaerobic chemotrophic alkaliphiles. In: J. Seckbach (Ed.) Journey to Diverse Microbial World. Kluwer. Dordrecht, The Netherlands. 191–208. Zhilina, T. N., G. A. Zavarzin, F. A. Rainey, E. F. Pikuta, G. A. Osipov, and N. A. Kostrikina. 1997. Desulfonatronovibrio hydrogenovorans gen. nov., sp. nov., an alkaliphilic sulfate reducing bacterium. Int. J. Syst. Evol. Microbiol. 47:144–149.

Prokaryotes (2006) 2:985–1011 DOI: 10.1007/0-387-30742-7_31

CHAPTER 1.31 ehT

s se l ro l oC

ruf l uS

a i re t caB

The Colorless Sulfur Bacteria LESLEY A. ROBERTSON AND J. GIJS KUENEN

The name “the colorless sulfur bacteria” has been used since the time of Winogradsky to designate prokaryotes that are either able, or believed to be able, to use reduced sulfur compounds (e.g., sulfide, sulfur and organic sulfides) as sources of energy for growth. Today, it is known that this group comprises a very heterogeneous collection of bacteria, many of which have little or no taxonomic relationship to each other. The colorless sulfur bacteria play an essential role in the oxidative side of the sulfur cycle (Fig. 1). Like all of the element cycles, the sulfur cycle has an oxidative and a reductive side, which, in most ecosystems, are in balance. However, this balance does not always exist, and accumulations of intermediates such as sulfur, iron sulfides, and hydrogen sulfide are often found. On the reductive side, sulfate (and sometimes elemental sulfur) functions as an electron acceptor in the metabolic pathways used by a wide range of anaerobic bacteria, leading to the production of sulfide. Conversely, on the oxidative side of the cycle, reduced sulfur compounds serve as electron donors for anaerobic, phototrophic bacteria or provide growth energy for the extremely diverse group of (generally) respiratory colorless sulfur bacteria. Common oxidation products of sulfide are elemental sulfur and sulfate (Fig. 1). The adjective “colorless” is used because of the lack of photopigments in these bacteria, although it should be noted that colonies and dense cultures can actually be pink or brown because of their high cytochrome content. This chapter will concentrate on the colorless sulfur bacteria, while the sulfate reducers and phototrophs will be discussed in 13 and 24. There is a wide range of different types of colorless sulfur bacteria with very diverse morphological, physiological and ecological properties and with equally diverse environmental requirements. Table 1 lists the genera that have traditionally been regarded as colorless sulfur bacteria (part A), as well as genera containing This chapter was taken unchanged from the second edition.

species originally not classified as such that have now been shown to be able to obtain energy from the oxidation of reduced sulphur compounds (part B). As will be discussed later, the apparent similarity of the metabolic pathways for sulfur oxidation disguises a high level of variation in these pathways indicating that the diversity among the colorless sulfur bacteria is probably due to convergent rather than divergent evolution. In addition to inorganic sulfur compounds, some species can also gain energy from the oxidation of other inorganic compounds such as hydrogen or ferrous iron. As well as differences in substrate range, there is also some variation in electron acceptor usage. Although most colorless sulfur-oxidizing bacteria require oxygen, a few are able to grow anaerobically using nitrogen oxides (e.g., nitrate) as their terminal electron acceptor during denitrification. One or two species (of the genus Acidianus) are capable of anaerobic metabolism by the reduction of sulfur (Segerer and Stetter, 1989), during which organic compounds or hydrogen serve as electron donors. Thiobacillus ferrooxidans is known to be able to reduce ferric iron under anaerobic conditions (Sugio et al., 1985). A somewhat exotic example of a sulfate reducer that might also be considered to be a colorless sulfur bacterium is Desulfovibrio sulfodismutans, which can grow anaerobically by the disproportionation of thiosulfate to sulfate and sulfide (Bak and Pfennig, 1987). Some of the reactions that generate energy from inorganic reduced sulfur compounds using oxygen and nitrate as electron acceptors are shown in Table 2. In the following sections, we will first discuss the physiology of the colorless sulfur bacteria, since physiology forms the basis of their present taxonomy, and then treat the taxonomy in the following section. This will be followed by a discussion of the habitats of the colorless sulfur bacteria, including artificial habitats, and finally some applications of their use. The chapter concludes with a brief section on the role of the colorless sulfur bacteria in the natural sulfur cycle, together with a description of the tech-

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CHAPTER 1.31 Fig. 1. The sulfur cycle. The colorless sulfur bacteria are involved primarily in those steps in which S2- and S are oxidized with O2 or NO3-. (Adapted from Bos and Kuenen, 1983).

organic sulfur corpounds assimilatory sulfate reduction

sulfate reserves (seawater)

mineralization processes

sulfidic minerals (e.g. pyrite)

dissimilatory sulfate reduction –

–

SO42

S2

biological oxidation with O2 or NO– 3

dissimilatory sulfur reduction

biological oxidation with O2 or NO– 3

anaerobic oxidation by phototrophic bacteria

spontaneous oxidation biological oxidation O2 or NO–3

S anaerobic oxidation by phototrophic bacteria sulfur deposits

Table 1. Genera of the colorless bacteria traditionally recognized as being capable of growth on reduced sulfur compounds and their environmental parameters. Anaerobic growth pH requirement Genus

Neutrophilic

Acidophilic

Thermal requirement Mesophilic

A. Traditional colorless sulfur bacteria Thiobacillus +a + Thiomicrospira + Thiosphaera + Sulfolobus + Acidianus + Thermothrix + Thiovulumd + Beggiatoa + Thiothrix + Thioplocad + Thiodendrond + Thiobacterium + Macromonas + Achromatiumd + + Thiospirad B. Other bacteria capable of growth on reduced sulfur Paracoccus + Hyphomicrobium + Alcaligenes + Pseudomonas + Hydrogenobacter + -

+ + + + + + + + + + + + compounds + + + + -

+, example known to exist; -, example unknown; V, variable. 16S rRNA analysis indicates a possible relationship. Hyperthermophilic archaebacterium. Axenic cultures are not available.

Denitrifier

S0/Fe3+ as electron acceptor

Symbiont

+ +c +c + -

+ + + + -

V + + + + + -

+ ?b -

+

+ + + -

-

-

Thermophilic

CHAPTER 1.31 Table 2. Examples of the reactions used by the colorless sulfur bacteria to gain energy for growth. H2S + 2O2 Æ H2SO4 2H2S + O2 Æ 2S0 + 2H2O 2S0 + 3O2 + 2H2O Æ 2H2SO4 Na2S2O3 + 2O2 + H2O Æ Na2SO4 + H2SO4 4Na2S2O3 + O2 + 2H2O Æ 2Na2S4O6 + 4NaOH 2Na2S4O6 + 7O2 + 6H2O Æ 2Na2SO4 + 6H2SO4 2KSCN + 4O2 + 4H2O Æ (NH4)2SO4 + K2SO4 + 2CO2 5H2S + 8KNO3 Æ 4K2SO4 + H2SO4 + 4N2 + 4H2O 5S0 + 6KNO3 + 2H2O Æ 3K2SO4 + 2H2SO4 + 3N2

niques available for the measurement of their activities.

Physiology The great diversity of colorless sulfur bacteria is also reflected in their physiology. This will come as no surprise if we remember that the group encompasses archaebacteria as well as eubacteria, and that the latter group is also very diverse, including common pseudomonads and organisms that might be considered “colorless blue green bacteria,” such as species of Beggiatoa. Most of our knowledge of the physiology of these organisms comes from the study of the relatively limited number of bacteria, such as the thiobacilli, that can be grown relatively easily in the laboratory. This is particularly true of our understanding of the biochemistry of sulfur metabolism and, to a lesser extent, of carbon metabolism. Although the biochemistry of the oxidation of sulfur compounds has received much attention over the last few decades, the pathways involved were not well understood. This was due, in particular, to the fact that the research was formulated around the hypothesis that there would be a single unifying enzymatic pathway for the oxidation of all reduced sulfur compounds. However, it is now clearly established that this is not the case. For example, the facultatively autotrophic Thiobacillus versutus and the obligately autotrophic T. tepidarius use two entirely different pathways (Fig. 2a and b). It should be noted that not only do the enzymes and electron carriers differ, but their localization in the membranes of the two species appears to be different. This is, of course, important for the mechanism behind the generation of a proton motive force (PMF) in these organisms. Little is known of the electron carriers involved in reverse electron transport for the production of reducing power during autotrophic growth, but all available evidence indicates that the PMF is the driving force for this process. For an extensive review of the state of the art, the reader is referred to Kelly (1988b).

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In most obligate and facultative autotrophs, the Calvin cycle serves as the route for carbon dioxide fixation. This is true, for example, for species from the genera Thiobacillus, Thiomicrospira, Thiosphaera, and Beggiatoa. Some other species, including those from Sulfolobus and Hydrogenobacter, possess a carbon dioxide fixation pathway based on a reductive Calvin cycle (Segerer and Stetter, 1989).

Energy and Carbon Sources or Electron Donors It has been common practice to subdivide the colorless sulfur bacteria in terms of their physiological type as defined mainly by their carbon metabolism. Table 3 defines these physiological types, which will be discussed briefly below. It should be remembered that some genera or species have not been studied in pure culture, and it is not yet known to which of the physiological groups they belong. Obligate Chemolithotrophs. These highly specialized bacteria require an inorganic source of energy and obtain their cell carbon from the fixation of carbon dioxide. As mentioned above, except in the case of the archaebacteria (which use a reductive carboxylic cycle [König and Stetter, 1989]), the colorless sulfur bacteria do this by means of the Calvin cycle (e.g., Schlegel, 1981). The citric acid cycle in these bacteria seems to be incomplete, and its enzymes probably serve a purely biosynthetic function. Despite their label as “obligate” autotrophs, it has been shown that many of these species actually can use small amounts of exogenous carbon compounds as a supplementary carbon source (Kuenen and Veldkamp, 1973; Matin, 1978), or can even ferment endogenous organic storage compounds such as glycogen (Beudeker et al., 1981; Kuenen and Beudeker, 1982), but these are both secondary metabolic activities, the organisms being primarily dependent on a lithotrophic energy source and carbon dioxide for autotrophic growth. Many Thiobacillus species, at least one species each from Sulfolobus and Hydrogenobacter, and all of the known species of Thiomicrospira fall into this group. Facultative Chemolithotrophs. These bacteria can grow either chemolithoautotrophically with an inorganic energy source and carbon dioxide, or heterotrophically with complex organic compounds providing both carbon and energy, or mixotrophically. Mixotrophy is the simultaneous use of two or more different metabolic pathways for energy and carbon (Gottschal and Kuenen, 1980). In the laboratory, mixotrophic growth is most easily observed

988

L.A. Robertson and J.G. Kuenen –S – SO– 3

SH2O Sulphite: cytochrome c oxidoreductase (M 44,000)

ENZYME A (M 16,000) Binds thiosulphate [A – SO3 – S –] in a 1:1 molar ratio

8H+ + 2O2

cytochrome oxidase ( 3)

Cytochrome M 260,000; aggregate of six polypeptides of M 43,000; 4–5 haem and 6–7 Fe per mole; Two mid-point redox potential centres, Em,7 –115 and +240 mV

ENZYME B (M 63,000; subunits.M 32,000; contains Mn in a 1:1 molar ratio

4H2O

8e–

cytochrome ss2

8e-

Cytochrome css2.5 M 56,000;subunits M 29,000; up to 3 haem and 3 Fe per mole; At least two mid-point redox potential centres,Em,7 – 215 and +220 mV

2–

2SO4 10H+

a

CHAPTER 1.31

Outer membrane

Cytoplasmic membrane

Periplasm

2–O3-S-SO3– trithionate trithionate hydrolase 4 H+

2 H2O

2–

2 SO4

2S2O32–

inhibition by FCCP

tetrathionate synthose S4O42-

S4O42-

–

2e

cytochrome oxidose

(Mechanism unkown) 2-

4 SO3

cytochrome c

8e-

Inhibition by HONO

4 H2O sulfite dehydrogenase 8 H+

Ubiquinone cytochrome b 4 SO24–

Outer membrane

Periplasmic space

4 SO24–

Cytoplasmic membrane

Cytoplasm

b

during continuous culture on limiting mixtures of substrates. The term mixotrophy usually designates simultaneous growth on a mixture of autotrophic and heterotrophic substrates (e.g., on thiosulfate and acetate). However, the simultaneous use of any mixture of substrates requir-

Fig. 2. Pathways of oxidation of reduced sulfur compounds in two different organisms. (a) The periplasmic thiosulfate-oxidizing system of Thiobacillus versutus as proposed by Kelly (1988a). The enzyme complex does not produce or metabolize polythionates such as tetrathionate. Thiosulfate is oxidized to sulfate without the formation of sulfur or other intermediates. Thiosulfate metabolism is initiated by its binding to enzyme A. In subsequent steps, sulfate is produced and released, while electrons are finally transferred to an aa3-type of cytochrome oxidase. (b) The periplasmic and cytoplasmic metabolism of trithionate, thiosulfate, and tetrathionate by Thiobacillus tepidarius as proposed by Kelly (1988b). In contrast to the system shown in part a, tetrathionate appears to be an intermediate in the oxidation of both thiosulfate and trithionate. After an initial hydrolysis of trithionate, yielding thiosulfate and sulfate, the thiosulfate is oxidized to tetrathionate. Available evidence indicates a periplasmic location of these systems. Tetrathionate is believed to be transported into the cell and then oxidized to sulfite in the cytoplasm by an unknown mechanism. Sulfite dehydrogenase is responsible for the final oxidation to sulfate, in which cytochrome b may be involved. FCCP, carbonylcyanide-p-trifluoromethoxyphenylhydrazone; HQNO, 2heptyl-4-quinolinol-1-oxide.

ing (partially) separate metabolic pathways or enzymes, and thus might produce diauxie or biphasic growth in batch culture (e.g., glucose and lactose, succinate and glucose, iron and sulfur, hydrogen and sulfide, acetate and lactate), could be considered as mixotrophy.

Table 3. Classification of the different physiological types of colorless sulfur bacteria.a Carbon source Physiological type Obligate chemolithotroph Facultative chemolithotroph (mixotroph) Chemolithoheterotroph Chemoorganoheterotroph (heterotroph)

Energy source

Inorganic

Organic

Inorganic

Organic

+b + -

+ + +

+ + + -

+ + +

Commonly used synonyms for chemolithotroph include chemolithoautotroph, autotroph, chemoautotroph, and lithotroph. +, used by the group; -, not used.

CHAPTER 1.31

The Colorless Sulfur Bacteria

Some of the thiobacilli, Thiosphaera pantotropha, Paracoccus denitrificans (Friedrich and Mitrenga, 1981), and certain Beggiatoa species (Nelson and Jannasch, 1983) are typical examples of organisms able to grow on mixtures of reduced sulfur compounds and organic substrates. To some extent, the phototrophic sulfuroxidizing bacteria might also be considered members of this group since most, if not all, of them are able to grow chemolithoautotrophically and mixotrophically on reduced sulfur compounds in the dark (Kuenen et al., 1985). Chemolithoheterotrophs. This little-known group of bacteria is characterized by an ability to generate energy from the oxidation of reduced sulfur compounds, but which cannot fix carbon dioxide. Until recently, Thiobacillus perometabolis was considered to be a member of this group, but it is now known that under certain conditions, it can grow autotrophically (KatayamaFujimura et al., 1984). However, unnamed chemolithoheterotrophic species have been isolated, and a few strains of Thiobacillus have been well characterized (Tuttle et al., 1974; Gommers and Kuenen, 1988). Some Beggiatoa strains may also belong in this group (Larkin and Strohl, 1983). As is clear from the example of T. perometabolis, careful testing under a variety of conditions is necessary in order to discriminate chemolithoheterotrophs from the facultative autotrophs as well as from the sulfur-oxidizing heterotrophs. Sulfur-Oxidizing Chemoorganoheterotrophs. Some heterotrophic bacteria can oxidize reduced sulfur compounds, but do not appear to derive energy from them. However, they may benefit from the reaction by the detoxification of metabolically produced hydrogen peroxide (e.g., some species of Beggiatoa, Macromonas, Thio-

_

o

989

+ pO (ell) 2 Tms. denitrificans T. denitrificans Tms. pelophila T. thioparus T. thiooxidans

Fig. 3. A “spectrum” showing the response of five different species of colorless sulfur bacteria to redox. The position of each line indicates the range of conditions of redox under which the organism can grow. T., Thiobacillus; Tms., Thiomicrospira. (Based on Timmer ten Hoor, 1977.)

bacterium, and Thiothrix) (Larkin and Strohl, 1983; Dubinina and Grabovich, 1984). The oxidation of thiosulfate to tetrathionate by many heterotrophic bacteria that do not seem to gain energy from the reaction is well documented (Tuttle and Jannasch, 1972; Tuttle et al., 1974; Mason and Kelly, 1988).

Electron Acceptors for Aerobic and Anaerobic Growth Oxygen is universally used among the colorless sulfur bacteria, although the degree of aerobiosis that can be tolerated by different species varies. The response of some of the colorless sulfur bacteria to redox can be demonstrated by means of a “spectrum” as shown in Fig. 3. Various colorless sulfur bacteria have different ways of growing or surviving anaerobically. One of the best studied is the use of nitrate or nitrite as a terminal electron acceptor, whereby the nitrogen oxides are reduced to nitrogen, a process termed denitrification. This will be discussed in detail in Chapter 23, but a brief consideration of the nitrate-reducing colorless sulfur bacteria is appropriate here. The denitrifying species tend to be neutrophilic (Table 4), but not necessarily mesophilic,

Table 4. Examples of the neutrophilic, mesophilic species capable of autotrophic growth on reduced sulfur compounds. Autotrophy Species Thiobacillus thioparus T. neapolitanus T. denitrificans T. novellus T. versutus T. intermedius T. perometabolis T. delicatus T. thyasiris Thiomicrospira pelophila Tms. denitrificans Tms. crunogena Thiosphaera pantotropha Beggiatoa sp. (marine) Beggiatoa sp. (freshwater) +, property present; -, property absent.

Denitrification

Obligate

Facultative

To NO2-

To N2

+a + + + + + -

+ + + + + + + + +

+ + + + + + + +

+ + + + + +

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L.A. Robertson and J.G. Kuenen

since at least one of the thermophiles (Thermothrix thiopara) can denitrify. A few (e.g., Thiobacillus thioparus) can only reduce nitrate to nitrite and require the presence of a nitrite-reducing bacterium for anaerobic growth (Table 4). Strictly speaking, of course, the latter reaction is not truly denitrification, but since the reaction still serves for electron acceptance and survival under anaerobic conditions, these species are appropriately included here. There are two known obligately chemolithotrophic sulfur bacteria that carry out complete denitrification to nitrogen. Thiobacillus denitrificans is relatively versatile in being able to grow under fully aerobic conditions with oxygen, and under fully anaerobic conditions with nitrate or nitrite (Aminuddin and Nicholas, 1973; Ishaque and Aleem, 1973). Thiomicrospira denitrificans is more fastidious. It grows well anaerobically with nitrate or nitrite, but can only use oxygen for growth if its concentration is kept extremely low (i.e., below the detection level of normal oxygen electrodes) (Timmer ten Hoor, 1975). These obligate autotrophs are far more efficient at anaerobic (denitrifying) growth on reduced sulfur compounds than the facultative species. Of the latter, only Thiosphaera pantotropha has been, thus far, found to retain its sulfuroxidizing potential under denitrifying conditions, but its mmax while doing so is extremely low (approx. 0.015 h-1) compared with those of Thiobacillus denitrificans and Thiomicrospira denitrificans (0.06 h-1). Other facultatively autotrophic bacteria lose their sulfur-oxidizing capacity in anaerobic cultures, but are still able to denitrify using organic compounds, or even hydrogen. Among these are Thiobacillus versutus and Paracoccus denitrificans (Taylor and Hoare, 1969; Friedrich and Mitrenga, 1981). Sulfide-dependent reduction of nitrate to N2 by Beggiatoa tufts has recently been shown using 15 N-labelled nitrate (Sweerts et al., 1990). Of the two sulfur-oxidizing genera of archaebacteria, Sulfolobus species appear to be the more dependent on oxygen, although some have been shown to use ferric iron and molybdate as electron acceptors under microaerobic conditions (Brock and Gustafson, 1976; reference is not an exact matchBrierly, 1982). Members of the genus Acidianus are able to grow under anaerobic conditions, by using hydrogen as the electron donor and sulfur as the acceptor, thus making these bacteria both sulfur-oxidizing and sulfur-reducing, depending on the conditions (Segerer and Stetter, 1989). Nelson and Castenholz (1981) have reported that some Beggiatoa species carry out an anaerobic reduction of intracellularly stored sulfur, using organic compounds such as acetate as electron donors. The ability of these organisms to oxidize sulfide

CHAPTER 1.31

to sulfur under aerobic conditions and then to reverse this reaction anaerobically would permit them to optimally profit from their habitat, where aerobic and anaerobic conditions frequently alternate. They may also actively migrate between aerobic and anaerobic zones. Even apparently obligately aerobic strains may have mechanisms allowing them to survive during anaerobiosis for a limited length of time. Thus, Thiobacillus neapolitanus, a species normally considered to be obligately respiratory, has been shown to be able to ferment internal reserves of polyglucose when confronted with anoxic conditions (Beudeker et al., 1981). As mentioned in the introduction, T. ferrooxidans can use ferric iron as an electron acceptor, although it is not yet clear whether this is linked to energy generation.

Ecophysiology as a Function of pH, Temperature and Nutrient Availability Colorless sulfur bacteria have been found growing at pH 9.0 and pH 1.0, at 4∞C and 95∞C, and at dissolved oxygen concentrations ranging from air-saturation to anaerobic levels (Table 1). It is obvious that a combination of physical, chemical, and (eco)physiological factors will suit the ecological niche of the organism within a particular microbial community. A number of these will be considered here. pH Range and Effects. The pH ranges of some of the colorless sulfur bacteria are surveyed in Table 1, and examples of neutrophilic and acidophilic species are listed in Tables 4 and 5. Within these ranges, of course, species often have different pH optima. The outcome of competition for a substrate at different pH values will therefore be dictated to a large extent by the pH optima of the competing bacteria. Thus, Kuenen et al. (1977) found that at pH values above 7.5, Thiomicrospira pelophila dominated

Table 5. Characteristics of acidophilic, mesophilic species capable of growth on reduced sulfur compounds and/or iron. Autotrophy Species Thiobacillus ferrooxidans T. thiooxidans T. albertis T. acidophilus Leptospirillum ferrooxidans

Utilization of

Obligate

Facultative

Sulfur

Iron

+

-

+

+

+ + +

+ -

+ + + -a

+

Also negative on other sulfur compounds, can use the iron in pyrite.

CHAPTER 1.31

thiosulfate-limited chemostat cultures, whereas when the pH was below 6.5, Thiobacillus thioparus was able to outcompete the other for thiosulfate. At intermediate pH values, the outcome of the experiments was not reproducible, with varying levels of the two populations. Apparently, the substrate affinities of the two species were so similar that other, less well-controlled variables (e.g., iron concentration, minor amounts of wall growth, etc.) became important for the outcome of the competition. Similar pH effects have been observed in the competition between T. versutus and T. neapolitanus (Smith and Kelly, 1979). The colorless sulfur bacteria that grow at neutral to slightly alkaline pH values are found in marine and freshwater sediments, soils, and wastewater treatment systems, to name but a few sources. As can be seen from Table 4, representatives of almost all of the genera fall within this group. Many of them have specialized in growth in the gradients where (anaerobic) sulfidecontaining zones come into contact with air or oxygen-containing water and will be discussed in the section on gradients. Some colorless sulfur bacteria are extreme acidophiles, able to grow at pH values as low as 1. As Table 5 shows, the group includes mesophilic obligate and facultative autotrophs (e.g., T. ferrooxidans and T. acidophilus, respectively). The acidophilic colorless sulfur bacteria are abundant in locations such as acid mine-drainage water, and it is therefore interesting that many of them are also able to oxidize (and gain energy from the oxidation of) metals such as iron. Thus, T. ferrooxidans is able to grow “mixotrophically” on the iron and sulfur components of pyrite (Arkestein, 1980) or on mixtures of ferrous iron and tetrathionate, gaining energy from the iron and sulfur oxidizing reactions (Hazeu et al., 1986, 1988). There have been a few reports of facultatively heterotrophic growth by T. ferrooxidans (e.g., Shafia and Wilkinson, 1969; Lundgren et al., 1964). However, it has since been shown that most of the T. ferrooxidans cultures available from culture collections were contaminated with acidophilic facultative autotrophs and heterotrophs (Harrison, 1984), including T. acidophilus and Acidiphilium cryptum, and it is now generally accepted that T. ferrooxidans is an obligate autotroph. It has frequently been assumed that T. ferrooxidans is one of the key species active in pyrite oxidation. In order to assess its likely significance for pyrite oxidation during coal desulfurization, Muyzer et al. (1987) used antibodies raised against T. ferrooxidans for an immunofluorescent assay of slurries made from coal from different sources. Unsterilized and sterilized coal samples were inoculated with T. ferrooxidans,

The Colorless Sulfur Bacteria

991

with a mixed culture of pyrite-oxidizing bacteria from a coal-washing installation, and a mixture of the two. Despite the fact that a DNAfluorescent stain indicated abundant microbial life in all of the slurries, the only sample in which a significant T. ferrooxidans population was detected was the control, which had been sterilized and then inoculated with the pure culture of T. ferrooxidans. It appears that in all other cases, other strains (which might include such species as T. thiooxidans, Leptospirillum ferrooxidans, or Acidiphilium cryptum, to name but a few) were able to successfully out-compete T. ferrooxidans for a niche in the consortium. Temperature As pointed out at the beginning of this section, colorless sulfur bacteria can be found growing at temperatures ranging from 4–95∞C. However, the majority of the wellstudied species are mesophilic. Although it is evident that the majority of natural environments are suitable for the growth of mesophiles, the diversity of the thermophilic organisms is likely to be much larger than suggested by Table 6, particularly in view of the recent discoveries of new thermophilic species among the colorless sulfur bacteria and other metabolic groups. Thus, it is clear that the species discussed in this section should be regarded as indicative rather than definitive. As most of the examples discussed elsewhere in this chapter will be taken from mesophilic bacteria, most of this section will be dedicated to consideration of the thermophiles. Thermophilic bacteria are generally associated with waters that have been geothermally heated. These range from warm springs, used for bathing since Roman times, through solfataras to submarine hydrothermal vents (e.g., Caldwell et al., 1976; le Roux et al., 1977; Jannasch, 1985). As can be seen from Table 6, the bacteria in this group can be subdivided into two groups, the moderate thermophiles (generally eubacteria), Table 6. Characteristics of moderately and extremely thermophilic species capable of growth on reduced sulfur compounds. Autotrophy Species Thiobacillus tepidarius T. aquaesulis Thermothrix thiopara Sulfolobus acidocaldarius Sulfolobus sp. HVS Acidianus infernus A. brierleyi

Obligate

Facultative

Temperature range (∞C)

+

-

20–52

-

+ +

30–55 72

-

+

60–85

+ + -

+

60–95 60–95 60–95

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which grow over the range 45–55∞C, and the extreme thermophiles (generally archaebacteria), some of which can grow at temperatures approaching 100∞C. Neutrophilic species make up the moderately thermophilic group. One neutrophile, Thermothrix (Tx.) thiopara has a higher optimum growth temperature (72∞C). This facultative autotroph was found in neutral (pH 7.0), hot (74∞C) springs (Caldwell et al., 1976; Brannan and Caldwell, 1980), where it forms macroscopic streamers as well as microscopic mats on the tufa. The streamers occur at the sulfide: oxygen interface (Caldwell et al., 1983), and the key role that oxygen plays in their development was demonstrated by means of a very simple experiment during which the surface of the hot spring was covered by a sheet of plastic to restrict entry of oxygen from the air. As a result of this, the dissolved oxygen dropped to 0.1 mg 1-1 from 3 mg 1-1, but other parameters such as pH and temperature were unaffected. The Tx. thiopara streamers then disappeared from their accustomed positions and reappeared at the edges of the sheet, where the sulfide:oxygen gradient had been reestablished. The acidophilic archaebacteria of the genera Sulfolobus and Acidianus represent the colorless sulfur bacteria among the hyperthermophiles. These genera include both obligately and facultatively autotrophic species. They are frequently found in association with sulfidic ores such as pyrite, chalcopyrite, and sphalerite. It has been suggested that the failure to find Sulfolobus species around hydrothermal vents, where Acidianus does occur, is due to the low salt tolerance of Sulfolobus species. Acidianus species can tolerate NaCl concentrations of up to 4% (Stetter, 1988). Of course, with growth temperatures between 60–95∞C, these strains seem almost “moderate” in comparison to the growth temperatures of the sulfur-reducing Pyrobaculum and Pyrodictium species (74– 110∞C). Nutrient Availability and Ecological Niches. Of the physiological types shown in Table 3, the obligate and facultative chemolithotrophs are the best known, having been the most extensively studied in pure and mixed cultures (e.g., Kelly and Kuenen, 1984; Kuenen, 1989; Kelly and Harrison, 1989; Kuenen et al., 1985; Kuenen and Robertson, 1989a, 1989b). One of the most important environmental parameters affecting the selection of these bacteria in freshwater environments was found by Gottschal and Kuenen (1980) to be the relative turnover rates of inorganic and organic components in the available substrates (Fig. 4). Thus, if the available substrate in energy-limited sys-

CHAPTER 1.31 H2S + 2O2 2H2S + O2 2S + 3O2 +2H2O Na2S2O3 + 2O2 + H2O 4Na2S2O3 + O2 + 2H2O 2Na2S4O6 + 7O2 + 6H2O 2KSCN + 4O2 + 4H2O 5H2S +8KNO3 5S + 6KNO3 + 2H2O

H2SO4 2S + 2H2O 2H2SO4 Na2SO4 + H2SO4 2Na2S4O6 + 4NaOH 2Na2SO4 + 6H2SO4 (NH4)2SO4 + K2SO4 + 2CO2 4K2SO4 +H2SO4 + 4N2 + 4H2O 3K2SO4 + 2H2SO4 + 3N2

Fig. 4. A model to describe the selection of different physiological types by the ratio of inorganic to organic substrates supplied in the medium. This model may also hold for complex (semi-natural) systems, where the relative turnover rates of the inorganic and organic compounds (or the ratio between the fluxes of these compounds) would determine the selection of different physiological types. For definitions of the various terms, see Table 3.

tems is wholly or predominantly inorganic, obligate autotrophs such as Thiobacillus neapolitanus will normally tend to dominate a community. Similarly, abundant organic substrates will generate communities dominated by heterotrophs. On mixed substrates, facultative autotrophs such as T. versutus or chemolithoheterotrophs will appear, depending on the ratio between the two types of substrate. If the substrate supply is predominantly organic, the sulfide-oxidizing heterotrophs or other heterotrophs will appear. This model was put to the test by means of a number of competition experiments in two- and three-membered mixed cultures of representatives from the physiological groups. In addition, a number of enrichment cultures inoculated from natural samples containing representatives of all of the physiological types were obtained. All of the experiments essentially showed that the predicted metabolic type became dominant (for example, see Fig. 5a and b). Although mathematical modelling predicted that in some cases pure cultures of only one metabolic type should be obtained, in practice, satellite populations of the others remained (Fig. 6). Clearly, secondary environmental or experimental conditions (e.g., excretion products such as glycollate, fluctuations in substrate or oxygen concentrations, and growth on the wall of the vessel) can result in deviations from the idealized model. It is obvious that a well-mixed chemostat is a model system that is rather remote from the common natural habitats of colorless sulfur bacteria, such as the sulfide:oxygen gradient in a sediment, and the results obtained can only demonstrate the principle. Moreover, the relative turnover rate of the organic and inorganic substrates is only one of the environmental parameters that determines the success of a particular species. Nevertheless, the use of this model (Fig. 4) has now clarified the situation, a practical consequence

CHAPTER 1.31

The Colorless Sulfur Bacteria 100

90

T.A2 (+ acetate)

80 70

T.A2 (+ glycollate)

60 50 40

T.neapolitanus (+ glycollate)

30

T.neapolitanus (+ acetate)

20

mixotroph

80 percentage of total cell-number

PERCENTAGE OF TOTAL CELL–NUMBER

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autotroph 20

10 0

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acetate (mM) 24 16 thiosulfate (mM) 2

16

20

8

0 3

PERCENTAGE OF TOTAL CELL–NUMBER

90

Fig. 6. Competition for acetate and thiosulfate in a chemostat between an autotroph, T. neapolitanus (open triangles); a mixotroph, T. versutus (closed circles); and a heterotroph, Spirillum G7 (open circles). The dotted lines indicate the results predicted from the model, the symbols indicate the actual results. The model held well for the extreme ratios of thiosulfate and acetate. However, although T. versutus dominated at intermediate ratios, as predicted, the other two types did not completely disappear. For the experimental details, see Fig. 5a. This model can be used for the selective enrichment of facultative autotrophs in chemostat cultures using an intermediate ratio of acetate and thiosulfate. (Based on Gottschal et al., 1982.)

80

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spirillum G 7

20 10 0

(b)

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THIOSULFATE (mM)

Fig. 5. The effect of organic or inorganic energy sources on competition. (a) The effect of different concentrations of organic substrates on the competition between Thiobacillus versutus (T. A2) and T. neapolitanus for growth-limiting thiosulfate in a continuous culture. The influent medium contained 40mM thiosulfate. During growth limitation by thiosulfate, it and the organic additives (where present) were used simultaneously by the mixed culture, and their actual concentrations in the chemostat were below the detection level. The graph shows the ratios of the two species at steady state. Open symbols, T. versutus; closed symbols, T. neapolitanus; circles, glycollate supplied; triangles, acetate supplied. (b) The effect of thiosulfate on the competition for acetate (10 mM) between T. versutus (T. A2) and a heterotrophic spirillum called G7. For experimental details, see (a). Open symbols, Spirillum G7; closed symbols, T. versutus. (Based on Gottschal et al., 1979.)

being that it has shown the way for the selective enrichment of facultatively autotrophic sulfur bacteria from fresh water. Steady-state conditions are more common in artificial environments than in nature, and therefore in order to test the effect of substrate fluctuations on the selection of the three representative species used in the experiments discussed above (Figs. 5a, 5b, and 6), Gottschal et al. (1981) ran chemostat cultures alternating feeds of acetate and thiosulfate. In twomembered cultures, the mixotrophic T. versutus was able to maintain itself on the substrate not used by whichever obligate species was involved, so that both species were subject to alternating periods of growth and starvation. However, in three-membered cultures, the two specialists were able to react more swiftly to the onset of substrate provision because of their constitutive enzymes, while the facultative species, which had to reinduce its autotrophic enzymes each time, disappeared. As with the steady-state experiments, when different mixtures of acetate and thiosulfate alternated, the outcome was deter-

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CHAPTER 1.31

0.4

T. thioparus 0.3

Tms. pelophila 0.2

0.1

Iron concentration Fig. 7. The specific growth rates (m) of Thiomicrospira pelophila and Thiobacillus thioparus as a function of the iron concentration in chemostat cultures at 25∞C. The graph was constructed from the results of competition experiments (at the growth rates indicated by the arrows at the y axis). The actual iron concentrations were not determined. (From Kuenen et al., 1977.)

mined by the concentrations involved. Enrichment cultures under this regime yielded a facultative autotroph that was able to avoid the need to induce its carbon dioxide fixation system by accumulating large amounts of PHB during the heterotrophic period. This work was carried out on aerobic, freshwater chemostat cultures and, as has been discussed in previous reviews (Kelly and Kuenen, 1984; Kuenen, 1989; Kelly and Harrison, 1989; Kuenen et al., 1985), marine enrichments are, for unknown reasons, generally less predictable. For example, mixotrophs did not form the dominant population in thiosulfate/acetate-limited marine cultures (Kuenen et al., 1985). That marine mixotrophs do exist has been shown by the isolation of a facultatively chemolithotrophic marine strain of T. intermedius from a thiosulfate-limited culture (Smith and Finazzo, 1981). Of course, factors other than the availability of electron donors can determine the type of population to be found in any given environment. For example, Kuenen et al. (1977) studied the effect of iron limitation and pH on the outcome of competition between two marine obligate autotrophs, Thiomicrospira (Tms.) pelophila and Thiobacillus (T.) thioparus. As can be seen from Fig. 7, Tms. pelophila will dominate mixed cultures of the two species at low iron concentrations, whereas T. thioparus will do better when iron is more abundant. One of the characteristics of Tms. pelophila is its tolerance of sulfide concentrations high enough to inhibit Thiobacillus spp. It has been postulated that sulfide inhibition is caused by the reaction of the sulfide with available iron, forming insoluble ferrous sulfide and thus drastically reducing the concentration of iron available for microbial

utilization. If this hypothesis is accurate, the ability of Tms. pelophila to grow well at very low iron concentrations would explain its sulfide tolerance.

Taxonomy Many of the colorless sulfur bacteria were discovered in the early years of microbiology, at a time when scientists were relying mainly on morphological characteristics to identify their organisms, and this fact is still reflected in our approach to their taxonomy. Needless to say, this has caused a certain amount of confusion (see Table 1 for an overview of the genera involved). The problems associated with the identification of some colorless sulfur bacteria have been aggravated because many of the bacteria involved are very specialized (e.g., obligate autotrophs) and, as a consequence, the number of physiological traits that can be screened is limited. This has resulted in relatively trivial features being given undue weight during classification. Taxonomy is a way of establishing identities and relationships in an attempt to create a sense of order among the various forms of life on earth. In ecology, as in other applications of taxonomy, the precise identification of a particular species may not always be as relevant as an accurate description of its physiological characteristics, but the comparison and correlation of data from different sources becomes easier if one can be certain, or even reasonably sure, of the identities of the various bacteria involved. Changes in taxonomic practice largely reflect new developments in available technology as well as improvements in our understanding of which factors indicate relationships, and which are merely resemblances. Taxonomic research into the colorless sulfur bacteria can thus be separated into three distinct, if overlapping phases, which will be discussed sequentially here.

Morphology The colorless sulfur bacteria, as a group, encompasses rods, spirals, cocci, filamentous cells and archaebacteria, and it comes as no surprise to find that the first of them to be described, Beggiatoa (Trevisan, 1842), is also one of the largest. The longest cells reported in the latest edition of Bergey’s Manual are 50 mm long (Strohl, 1989), but a recent paper described the observation of a marine strain more than 100 mm long (Nelson et al., 1989). Another morphologically distinct genus, Thiothrix, was described by Winogradsky in 1888, but it was not until 1904 that Beijerinck described the first of the smaller colorless sulfur bacteria, Thiobacillus thioparus.

CHAPTER 1.31

As may be seen from a survey of the relevant chapters in Bergey’s Manual a few genera are still, today, based largely on morphological descriptions (e.g., Thiospira, Macromonas, Thiovulum) because pure cultures are either not available, or have only recently been achieved. In addition to cell size and shape, other morphological details that have been considered important are the appearance of inclusion bodies such as sulfur or poly b-hydroxybutyrate (PHB), number and placement of flagella, colony size, colony form and colony color. One of the dangers associated with too strong a reliance on such features is that all of them can vary depending on the growth conditions. As a single example of this problem, the faculatively autotrophic Thiosphaera pantotropha might be considered. When grown autotrophically on thiosulfate, it occurs as small cocci (0.7 ¥ 0.9 mm), which are generally found singly or in pairs (Fig. 8a). Cultivation in batch culture on rich media in which rapid growth will occur leads to a slightly larger, pleomorphic form (Fig. 8b). In chemostat cultures on mineral medium with acetate, chains of cocci appear. The internal structure of Thiosphaera pantotropha also changes with its growth conditions. Thus the normal appearance, with few inclusions, of a Gram-negative organism, which is found during substrate-limited chemostat culture (Fig. 8c), gives way to cells with PHB granules and complex membranous structures (Fig. 8d) when grown under oxygen or nitrogen-limited conditions, or in the presence of hydroxylamine. Cultivation on acetone or propan-2-ol results in the formation of large, crystalline structures (Fig. 8e), while denitrifying growth on sulfide can result in the accumulation of a fine deposit of sulfur in the periplasm (Fig. 8f). The colonial form of this species also varies, with off-white, translucent colonies being produced during growth on mineral medium with acetate or thiosulfate; and larger, thicker, browner colonies being generated during growth on rich media. Even the obligate autotrophs, which with their more limited range of growth conditions might appear to have less scope for variation, can produce substantial morphological changes. Thus, the number of carboxysomes formed by Thiobacillus neapolitanus increases dramatically under CO2 limitation (Beudeker et al., 1980), and polyglucose inclusions appear under nitrogen limitation (Beudeker et al., 1981). From all of this, it is clear that while valuable information can be gained from morphological studies on cells or colonies grown under welldefined conditions, this information should be used cautiously and in conjunction with other data. In exceptional circumstances, very distinctive morphology (e.g., in the case of Beggiatoa or

The Colorless Sulfur Bacteria

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Hyphomicrobium) might be more reliable as an indicator of identity.

Physiological Screening As more pure cultures became available, it became possible to determine the physiological capabilities of different bacteria, and physiological criteria gradually became an integral part of the taxonomists’ armory. For the obligate autotrophs, these might include such tests as optimum pH, growth temperature, ability to denitrify, and (generally very limited) substrate range. In addition to these, the facultative autotrophs are generally subjected to the same range of tests used for heterotrophic bacteria including oxidase, catalase and urease reactions, and the ability to grow on or generate acid from a range of substrates. An extensive study of the Thiobacillus species then available resulted in a numerical taxonomy analysis of the genus (Hutchinson et al., 1969) that recognized that “species” such as Ferrobacillus ferrooxidans and Thiobacillus thiocyanoxidans were actually strains of existing species (T. ferrooxidans and T. thioparus, respectively). The tests recommended by Hutchinson et al. (1969) for the identification of new Thiobacillus species included growth on sulfide, sulfur, thiocyanate, citrate and nutrient broth, the amount of thiosulfate used, sulfur deposition, and the effect of inhibitors such as streptomycin, bacitracin and ampicillin. In many respects, the range of substrates on which an isolate is tested is defined by the interests of the research group. The reduced sulfur compounds are not included in standard test batteries, and the sulfur-oxidizing abilities of many bacteria are only now being discovered. For example, Friedrich and Mitrenga (1981) tested a number of hydrogen-oxidizing bacteria and found that many of them, including Paracoccus denitrificans and some Alcaligenes species, were able to grow autotrophically on thiosulfate. Attempts to use thiosulfate as an inhibitor of heterotrophic nitrification by a “Pseudomonas” species gave anomalous results until it was realized that the culture was growing mixotrophically, using both the acetate supplied as the primary growth substrate and the thiosulfate added as a possible inhibitor. Subsequent experiments revealed that this “Pseudomonas” species was also able to grow autotrophically using reduced sulfur compounds (Robertson et al., 1989). A problem associated with the use of substrate ranges for taxonomic purposes is that it is difficult to determine how closely related bacteria with the same enzyme system are. Thus, possession of the Calvin cycle enzymes for carbon dioxide fixation or the denitrification pathway enzymes is not considered sufficient grounds for

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CHAPTER 1.31

a

b 5

c

e

d

f

Fig. 8. Variations in the morphology of cells of Thiosphaera pantotropha in relation to growth conditions or substrates as seen under the electron microscope. (a) Aerobic, autotrophic growth on thiosulfate, Pt shadowed. (b) Aerobic, heterotrophic growth on a mixture of acetate, fructose, and yeast extract, Pt shadowed. (c) Thin section of cells from an acetate-limited, chemostat-grown culture, stained with ruthenium red to show the membrane structures. (d) Thin section of a cell from an aerobic, acetate-limited chemostat with hydroxylamine, stained with ruthenium red to show the membrane structures. The white bodies are PHB granules. (e) Thin section of an acetone-grown cell showing crystalline inclusions. (f) Thin section of an anaerobic (denitrifying) cell grown on sulfide and stained with silver to show the periplasmic deposits of sulfur. (Fig. 8b from Robertson and Kuenen, 1983b. Fig. 8c from Bonnet-Smits et al., 1988. Fig. 8f, courtesy of H. J. Nanninga. All electron microscopy courtesy of W. Batenberg.) All bars = 0.5 mm.

CHAPTER 1.31

The Colorless Sulfur Bacteria

classifying the relevant bacteria into a single group, and it must be questioned whether the sulfur-oxidizing enzymes are a better indicator, especially since there appears to be several different pathways involved (Kelly, 1988a, 1988b) (see also Fig. 2, above). Certainly, it is recognized that at least one genus, Thiobacillus, is very heterogeneous (Kuenen, 1989; Kelly and Harrison, 1989) and will probably require subdivision. It has been suggested that this separation should be made between the obligate and the facultative autotrophs (thus, again on physiological grounds) but, as will be seen in the following section, this is probably not sufficient.

Analytical Techniques The determination of the GC content of the DNA of bacterial isolates has been used for a long time to determine whether or not strains could be related. It is, to some extent, a negative test because, while widely differing GC values could confirm that two strains were not related, matching GC values do not guarantee that they are the same. Cellular fatty acid analysis has been used in the taxonomy of the Thiobacilli (Agate and Vishniac, 1973; Katayama-Fujimura et al., 1982). Katayama-Fujimura et al. (1982) also included the analysis of ubiquinones and DNA base composition in their study. They initially subdivided the bacteria into groups based on whether they were obligately or facultatively autotrophic, and then on the basis of their possession of menaquinone 8 or 10, (MK-8 or MK-10) and then used the fatty acid analysis to further examine each group. This led to a proposal for the grouping of the different strains, which is shown in Table 7. Some of the first publications to consider the Thiobacilli in relation to other colorless sulfur bacteria involved the phylogenetic analysis of the various species by comparison of their 5S

997

rRNA sequences (Lane et al., 1985; Stahl et al., 1987). This work has now been extended by the use of 16S rRNA analysis (Lane et al., 1990; Oyaizu et al., 1990), and has revealed that there are closer matches between some sulfuroxidizing bacteria and other apparently unrelated strains such as Escherichia coli than between these and other sulfur oxidizers. Table 8 summarizes some of the results from the 5S and 16S rRNA comparisons. The sulfur oxidizing genera Sulfolobus and Acidianus are archaebacteria and therefore not listed in Table 8. If the initial separation into obligate and facultative autotrophs employed by KatayamaFujimura et al. (1982) is removed, it can be seen that the results in Tables 7 and 8 support each other. Thus groups I.1 and I.2 from the menaquinone/fatty acid analysis correspond to group alpha from the 16S rRNA, groups II and III-1 with group beta-1, and groups III-2 and III3 with beta-2. Of course, the range of bacteria subjected to the menaquinone/fatty analysis was much more limited than that in the 5S and 16S rRNA survey, and more data would be useful. However, such independent agreement must confer additional weight that chemotaxonomy and phylogeny may provide more reliable tools for the classification of these bacteria than physiological or morphological observations.

Habitats As may be deduced from the range of physiological characteristics discussed above, the colorless sulfur bacteria, in one form or another, are to be found in almost every life-supporting environment where reduced sulfur compounds are found. Because the range of habitats is so wide, the principles underlying the selection of colorless sulfur bacteria in selected situations will be be discussed below. The following section will then deal more generally with the role of the

Table 7. Classification of the Thiobacillus species based on analysis of their menaquinone and fatty acid composition. Autotrophy type Facultative Facultative Facultative Facultative Facultative Facultative Obligate Obligate Obligate Obligate Obligate

Menaquinone

Hydroxy fatty acid

Species

Group

MK-10 MK-10 MK-10 MK-8 MK-8 MK-8 MK-8 MK-8 MK-8 MK-8 MK-8

None 3OH 10:0 3OH 14:0 3OH 10:0 3OH 10:0, 3OH 12:0 3OH 10:0, 3OH 12:0 3OH 10:0, 3OH 12:0 3OH 10:0, 3OH 12:0 3OH 12:0 3OH 14:0 3OH 14:0

T. novellus T. versutus T. acidophilus T. delicatus T. perometabolis T. intermedius T. denitrificans T. thioparus T. neapolitanus T. ferrooxidans T. thiooxidans

I.1 I.1 I.2 II II II III.1 III.1 III.2 III.3 III.3

MK, menaquinone. The number indicates the number of isoprenoid units. Groupings are as proposed by Katayama-Fujimura et al. (1982).

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CHAPTER 1.31

Table 8. Classification of the colorless sulfur bacteria and examples of apparently related species (group “purple”), also termed Proteobacteria (Stackebrandt et al., 1988), as shown by 16S rRNA analysis.a Main group Alpha

Beta

Borderline Gamma

Delta

Subgroup

Species

1 1 2 2 1 1 1 1 2 2

Thiobacillus (T.) acidophilus, Acidiphilium rubrum A. cryptum, T. novellus Rhodobacter capsulatus, T. versutus Paracoccus denitrificans T. denitrificans, T. thioparus T. intermedius, T. perometabolis Rhodocyclus gelatinosa Vitreoscilla T. tepidarius, T. ferrooxidans T. albertis, T. thiooxidans T. neapolitanus, Chromatium vinosum Thiothrix nivea, Riftia symbionts Thiomicrospira pelophila, Thiomicrospira L-12 Bathymodicius symbionts Other symbionts Pseudomonas aeruginosa, P. putida Beggiatoa alba, Beggiatoa sp. Escherichia coli, Salmonella, Proteus, Vibrio Thiovulum, Campylobacter, Wollinella

1 1 1 1 1 1 2

Atypical strains have been omitted for the sake of simplicity. Adapted from Lane et al., 1990; and Harrison (1989).

colorless sulfur-oxidizing bacteria in the sulfur cycle, and this discussion of habitats is not intended to be exhaustive. In natural habitats, the reduced sulfur compounds available tend to be either sulfides (including metallic ores) or sulfur. Thanks to the activities of sulfate-reducing bacteria, especially in anoxic sediments, hydrogen sulfide is very commonly available, and some algal and cyanobacterial mats have been shown to generate organic sulphides (e.g., Andreae and Barnard, 1984). One of the main factors that bacteria growing on hydrogen sulfide have to contend with is the chemical reaction between sulfide and oxygen, and therefore the colorless sulfur bacteria are frequently found in the gradients at the interface between anoxic, sulfide-containing areas and aerobic waters and sediments where, at very low oxygen and sulfide concentrations, they can effectively compete with the spontaneous chemical oxidation reaction. Of course, the rate of chemical oxidation of metal sulfides with oxygen is very low at acid pH levels, so that the acidophilic bacteria need not, therefore, occur predominantly in gradients, as their neutrophilic counterparts must. The same holds for deposits of elemental sulfur, which does not react spontaneously with oxygen at a significant rate. Another habitat in which sulfide-oxidizing bacteria appear to be of some importance is in the complex communities of prokaryotes and eukaryotes around hydrothermal vents, where the sulfide is geologically rather than biologically generated. In the course of research into the life around these vents, it was shown that many

invertebrates have symbiotic colorless sulfur bacteria, and this can itself be regarded as a distinct habitat (Cavanaugh et al., 1981). A third example of a type of habitat for these bacteria that is becoming steadily more common is that associated with human activities, largely in connection with waste treatment and industrial leaching of ores for (heavy) metal recovery.

Gradients in Aquatic Systems and Sediments Sulfide:oxygen gradients occur in stratified water bodies, as well as in soils and sediments. Such gradients can range in size from a few hundredmicrometers-thick in a microbial mat or surface sediment to several meters in a stratified body of water (Sorokin, 1970, 1972; Jørgensen et al., 1979). These gradients can sometimes be distinguished with the naked eye. For example, Thiovulum grows as a fine white veil at the interface between sulfide and oxygen (Jørgensen, 1988). Wirsen and Jannasch (1978), studying the effect of the sulfide:oxygen gradient on the formation of these veils in continuous flow cultures, observed that the veils dispersed within minutes of the cessation of the flow of sea water through the culture vessel, and formed again once the flow was resumed, indicating chemotaxis of the swarming form of Thiovulum toward critical concentrations of oxygen and sulfide. The genus Beggiatoa contains marine and freshwater species that are typical of life at the aerobic:anaerobic interface. Dense mats of

CHAPTER 1.31

almost axenic cultures of Beggiatoa on sulfidecontaining sediments are frequently observed, especially in marine sediments where sulfide production rates can be very high. These mats are characterized by very steep oxygen and sulfide gradients over a few mm (Jørgensen, 1982, 1988). Since Beggiatoa oxidizes the sulfide at a very high rate, the overlying aerobic water is effectively “protected” from diffusion of toxic sulfide. The typical conditions for growth in this type of mat have been very difficult to reproduce in the laboratory. Indeed, they are so specialized that it was only recently, when available techniques had improved sufficiently to allow in vitro cultivation on sulfide:oxygen gradients, that the autotrophic potential of marine strains was established unambiguously (Nelson and Jannasch, 1983; Nelson, 1988) (see also Chapter 166). The Beggiatoa cells were cultured in closed tubes using a layer of very soft (0.2%) agar over a sulfide-containing plug of harder (1.5%) agar, thus allowing the formation of an upward sulfide gradient. Diffusion from a headspace containing air contributed a downward oxygen gradient. The Beggiatoa colony grew as a “plate” that was less than 1 mm thick at the point where the two gradients overlapped. The very rapid oxidation of sulfide allowed the organisms to maintain an extremely low concentration of the two substrates. As a result, chemical oxidation of sulfide was insignificant. For example, the turnover time for sulfide and oxygen was only 3 seconds in Beggiatoa gradients, whereas the half life of these two substances in sterile controls was about 20 min. Enzyme analysis and the fixation of 14CO2 by these cells confirmed that they were capable of autotrophic growth. The situation regarding freshwater strains is not so clearcut. Schmidt et al. (1987) showed sulfide oxidation rates for a freshwater strain comparable to those obtained with the marine strain discussed above, but further experimentation is necessary in order to establish whether energy for growth can be derived from the reaction. Another well-known place where gradients occur is within phototrophic mats. Jørgensen and des Marais (1986) studied the zonation around a cyanobacterial mat growing in a hypersaline pond and found that a band of Beggiatoa occurred 1.5 mm below the cyanobacteria. The photosynthetic activity of the cyanobacteria generated sufficient oxygen to produce an oxygen peak with a maximum of 1mM at the cyanobacterial band. A steep downward gradient of oxygen overlapped a sulfide gradient at the point where the Beggiatoa were growing. In an earlier study, Jørgensen (1982) described the diurnal changes in the sulfide and oxygen gradients and the microbial community to be found in a sulfuretum (a microbial mat in which the total turn-

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over of inorganic and organic compounds is heavily dominated by the sulfur cycle) on the surface of a sediment. It was observed that the mixture of cyanobacteria, phototrophic sulfur bacteria, and Beggiatoa was stratified, and that the relative positions of the the three populations among the strata were governed by the level of photosynthetically generated oxygen (Fig. 9). During the night, when the oxygen had been depleted and the oxygen boundary extended to the surface of the sediment, the phototrophic Chromatium was found at the surface. However, once photosynthesis began, with the onset of daylight, oxygen began to build up in the sediment, and the Chromatium followed the sulfide boundary down, remaining within the anaerobic part of the sediment. The Beggiatoa population tended to move with the sulfide:oxygen interface, except during the night when this was in the stagnant water above the surface of the sediment. As Beggiatoa is only motile by means of a gliding action, it is restricted to the solid phase. Other conspicuous colorless sulfur bacteria such as Thiothrix, Thioploca, and Archromatium have all been encountered as typical organisms in such gradients. Furthermore, mixed cultures of Thiobacillus-like bacteria sampled from sulfide:oxygen gradients and showing active sulfidedependent carbon dioxide fixation clearly exhibit chemotaxis toward the interface when transferred to artificial sulfide:oxygen gradients in the laboratory (J. G. Kuenen, unpublished observations).

Hydrothermal Vents An interesting extension of the model for the selection of freshwater colorless sulfur bacteria discussed above is to be found in the results of research on the mesophilic bacterial communities found around the different hydrothermal vents (Jannasch, 1985, 1988). These vents are a result of the movements of the tectonic plates of the earth’s crust. Seawater penetrates deep under the sea floor and is heated geothermally, reaching temperatures as high as 1,200∞C. Under these conditions, it reacts with and dissolves various reduced chemicals before being forced to the surface again as hydrothermal fluid, which contains sulfide, CO2, and methane, as well as various metals and hydrogen. The type of vent that occurs depends very much on the overlying geology, and can be at least partially separated into “bare lava” and “warm” systems. In the bare lava vents, the pressurized hydrothermal fluid reaches the surface of the sea floor at temperatures around 350∞C. As it issues from the vents, it reacts with chemicals in the sea water, forming precipitates that often accumulate as “chim-

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CHAPTER 1.31

distance/mm

–1

O2

0

Oscillatoria Beggiatoa

1 2

Chromatium

H2S 12

distance/mm

–1

09

O2

15 O2

day

0

Beggiatoa 1 2

06 H2S

18

Oscillatoria

Beggiatoa night

Chromatium 03

Oscillatoria 21

H2S

Chromatium

24

distance/mm

–1

O2

Chromatium

0

Beggiatoa 1

H2S

2

Oscillatoria Chromatium

Fig. 9. Diurnal cycle of oxygen and sulfide distribution and of microbial zonation in a marine sulfuretum. The zero line in each box indicates the interface between the sediment and the overlaying water phase. The dominant genera at each stratum are indicated in each box. Diatoms were primarily seen among the Oscillatoria. In addition to diurnal changes in light, oxygen, and sulfide, another important factor was that the Beggiatoa which are gliding bacteria could not move out of the sediment, whereas Chromatium, which is also motile, was able to move into the water phase above. From Jørgensen (1982).

neys.” Because the formation of metal sulfides gives the fluid issuing from these chimneys the appearance of smoke, they have become known as “black smokers.” The “warm” vents, on the other hand, are the result of the hydrothermal fluid percolating through sediments on its way to the surface, and the solution tends to be much cooler (